Organ/tissue decellularization, framework maintenance and recellularization

ABSTRACT

Methods for decellularizing organs and tissues in vitro and in vivo are provided, as are methods of maintaining organ and tissue frameworks and methods of recellularizing organs and tissues, thereby providing an approach to needed organs or tissues.

FIELD

The disclosure relates generally to the decellularization of diseased orinjured organs or tissues and to the recellularization thereof.

BACKGROUND

In the United States, more than 2,600 kidneys are discarded annually,from the total number of kidneys procured for transplant. In organbioengineering and regeneration, seeding of cells on supportingscaffolding material offers a potential route to clinical application[1-24]. Discarded organs represent a catastrophic and unrelenting figureat a time of dramatic organ shortage and urgent need for new,potentially inexhaustible sources of transplantable organs.

Cell-scaffold technology has allowed researchers to manufacturerelatively simple structures such as vessels, bladders, segments ofupper airways and urethras from the patients' own cells, which wereeventually implanted in patients with good short- and mid-term results.If we exclude the case of bioengineered vessels, the new structures wereimplanted without being reconnected to the vascular system, on theassumption that tissues of the bioengineered structures would initiallyreceive oxygen and nutrients by diffusion from adjacent vascularisedtissues of the host until neoangiogenesis occurred. While this strategy(i.e., not reconnecting the new structure to the recipient'svasculature) may be acceptable for thin structures such as the onesmentioned above, it is inadequate for more complex, “thicker” organslike the kidney, which requires reconnection to the patient's vascularsystem to maintain cell viability and the core of the organ, as well as,to exert function.

Organ transplantation is one of the greatest achievements in the historyof modern medicine. In view of the excellent results attained to date,the demand for transplantable organs is escalating, whereas the supplyhas reached a plateau. Consequently, waiting times and patients'mortality while on the waiting list are increasing dramatically (FIG.1A) and so the identification of new, potentially inexhaustible sourcesof organs has become an urgent need.

SUMMARY

The disclosure provides materials and methods that diminish theaforementioned need by a new source of tissues and organs claimed fromtissues and organs previously discarded. The methods provide an approachto the decellularization of tissues and organs that are diseased orotherwise deemed unusable in transplant scenarios. Followingdecellularization, disclosed herein are methods for maintaining theacellular tissue or organ scaffold or framework composed largely ofextracellular matrix with intact vasculature. Also provided are methodsfor recellularizing the tissue or organ scaffold ex vivo or in vivo,thereby regenerating a healthy tissue or organ.

In one aspect, the disclosure provides a method of regenerating a tissueor organ comprising inserting an acellular tissue or organ scaffold intoa subject in need thereof, establishing at least two vascularconnections of the scaffold vasculature and the subject vasculature, andincubating the acellular scaffold until an intact tissue or organ isproduced. It is expected that this method will prove advantageous fortissues and organs too thick to obtain sufficient in vivo oxygen andnutrients via diffusion to be viable and thrive. In particular, thedisclosure provides a method of regenerating a tissue or organ for asubject in need comprising: (a) providing an acellular tissue or organscaffold; (b) contacting the tissue or organ scaffold with a cellcharacteristic of the tissue or organ; (c) incubating the scaffold andcell ex vivo under static conditions; (d) implanting the incubatedscaffold and cell into the subject and establishing at least twovascular connections of the scaffold vasculature and the subjectvasculature; and (e) incubating the acellular scaffold until an intacttissue or organ is produced. In some embodiments the cell is obtainedfrom the subject. In some embodiments, the scaffold is contacted with atleast 5×10⁷ cells. An example of a cell for use in the method is avascular endothelial cell.

A variation of the method further comprises incubating ex vivo thescaffold and cell under ramping perfusion conditions, e.g., wherein theperfusion comprises delivery of at least 1×10⁸ cells at a varying ratespanning 2 ml/minute to 20 ml/minute. Some embodiments of any of theabove methods further comprise attachment to the scaffold of an antibodyproduct specifically binding to the cell, such as an antibody productcomprises an antigen binding site for CD31. By “antibody product” ismeant a complete antibody or any fragment or variant form thereof thatretains at least the six complementarity determining regions specifyingthe binding specificity of the antibody product. In some embodiments,the method further comprises attachment of an anti-thromboticcomposition to the scaffold, such as attachment of heparin to thescaffold. In various embodiments, the organ is a kidney, pancreas,liver, gall bladder, stomach, small intestine and large intestine;exemplary organs include a kidney, a pancreas or a liver.

In another aspect, the disclosure provides a method of decellularizing atissue or organ comprising: (a) perfusing an ex vivo tissue or organwith at least 50 volumes of a detergent; and (b) rinsing the ex vivotissue or organ with at least 50 volumes of a neutral buffer. Thedetergent may be either ionic or non-ionic. In some embodiments, theorgan is selected from the group consisting of kidney, pancreas, liver,gall bladder, stomach, small intestine and large intestine, for examplekidney, pancreas or liver. For kidney decellularization and thedecellularization of other hardy organs, an ionic detergent, e.g.,sodium dodecyl sulfate, is used. For less hardy organs such as thepancreas, a non-ionic detergent, e.g., Triton X-100, is used. Someembodiments comprise a method wherein the perfusing and rinsing areperformed using at least two fluid inlets and at least one fluid outlet.The methods may be performed using a variety of fluid flow rates, aswould be understood in the art. In some embodiments of the method, theflow rate of the two fluid inlets is about equal. In some embodiment,the flow rate of the fluid outlet is at least 5 ml per minute.

The method described above may further comprise an initial step ofdelivering distilled water to the organ or tissue at a rate of about 12ml per minute for at least 10 hours. Some applications of the methodfurther comprise delivering at least one liter of 0.0025 w/w % DNasesolution to the detergent-treated tissue or organ prior to rinsing thetissue or organ.

The method described above contemplates embodiments wherein the ionicdetergent is an anionic detergent. In some embodiments, the anionicdetergent is an organosulfate detergent, e.g., sodium dodecyl sulfate.For example, the disclosure contemplates embodiments wherein the sodiumdodecyl sulfate is 0.5% sodium dodecyl sulfate. As noted above, otherembodiments use a non-ionic detergent such as Triton X-100.

Some embodiments comprise a method wherein the tissue or organ isperfused with at least 30 liters of ionic detergent. Some embodimentscomprise a method wherein the perfused tissue or organ is rinsed with atleast 40 liters of neutral buffer. Also comprehended are methods whereinthe perfusing step is performed for at least 10 hours.

In some embodiments, the neutral buffer is phosphate-buffered saline.Further, some embodiments are contemplated wherein the rinsing step isperformed for at least 40 hours. Some embodiments comprise a methodwherein the neutral buffer contains 10 U/ml heparin.

Further comprehended are embodiments wherein the method furthercomprises exposing the tissue or organ to a bactericidal agent. Anybactericidal, microbicidal, bacteriostatic, or anti-microbic agent knownin the art to be compatible with eukaryotic cells, tissues and organs,such as human cells, organs and tissues, is contemplated by thedisclosure, including but not limited to betadine.

Other features and advantages of the disclosure will become apparentfrom the following detailed description, including the drawing. Itshould be understood, however, that the detailed description and thespecific examples, while indicating preferred embodiments, are providedfor illustration only, because various changes and modifications withinthe spirit and scope of the invention will become apparent to thoseskilled in the art from the detailed description.

BRIEF DESCRIPTION OF THE DRAWING

FIG. 1. Statistical data on kidney transplantation. A) United Networkfor Organ Sharing data on the number of patients who deceased whilewaiting for kidney transplantation. This number has dramaticallyincreased in the last decade due to longer waiting times determined bythe fact that the constantly growing number of patients registered inthe waiting list is not being balanced by an adequate offer oftransplantable kidneys. B) United Network for Organ Sharing data onhuman kidneys discarded from transplantation. Basically, one fifth ofthe procured kidneys are discarded as not suitable for transplantationfor multiple reasons.

FIG. 2. Histological examination of decellularized human kidneys. H&E,Masson trichrome, methenamine silver and DAPI staining of native (A, C,E, G, I) and acellular (B, D, F, H, J) human kidneys. Cell and nuclearmaterial are well cleared as demonstrated by the total lack of nucleiand cellular components or debris. The inset in B shows an example of aseverely sclerotic glomerulus that is now completely acellular. Arrow inpanel J shows the intact framework of a glomerulus.

FIG. 3. Removal of antigenic markers. Immunostaining for HLA ABC (classI, A, B), HLA DR (class II, C, D) antigens and integrin a3b1 (E, F) innative (A, C, E) and acellular (B, D, F) human kidneys. The staining ofthese antigenic markers is completely absent in the acellular ECMscaffolds.

FIG. 4. Preservation of ECM molecules. Immunostaining for Collagen typeIV (AeD), laminin alpha5 (EeF), agrin (GeH) and nidogen (IeJ) in native(A, C, E, G, I) and acellular (B, D, F, H, J) human kidneys. The naturallocation of these ECM molecules is shown in the native kidneys. There iswell-preserved expression of the ECM molecules following thedecellularization process, and they remained in their natural locations.

FIG. 5. Infrastructural analysis. Scanning electron microscope images ofnative (A, B) and acellular (C, D, E, F) human kidneys. The imagesindicate good preservation of the ultrastructure in the acellular ECMscaffolds, compared with the native kidneys. No cellular remnants can bedetected on the acellular matrix.

FIG. 6. Angiogenic induction. The angiogenic potential of the renal ECMscaffolds was tested using CAM assay. Images of the scaffolds placed onthe chorioallantoic membrane from day 0, 5, and 7 (B, C and D,respectively) show progressive attraction of blood vessels towards theECM scaffold. The number of vessels induced by the implantation of ECMversus control (polystyrene membranes) was calculated from these images(A).

FIG. 7. Resistance to vascular pressures. Pressures within the renalartery in acellular human renal ECM scaffolds increased linearly withflow rate, similar to native kidneys. Linear regression calculationshows similar trends, which however significantly differ between nativeand acellular kidneys.

FIG. 8. Hemotoxylin & Eosin staining of kidney tissuepost-decellularization obtained by arterial perfusion only. Cellclearance is not complete, as revealed by the presence of cells andnuclei detected in glomeruli as well as in other zones of the kidneyparenchyma.

FIG. 9. DAPI staining of kidney tissue post-decellularization obtainedby arterial perfusion reveals the presence of cells, as indicated byfluorescence (bright dots) staining of cell nuclei.

FIG. 10. Schematic illustration of organ or tissue decellularization.

FIG. 11. Pancreas decellularization through three different inlets,i.e., the pancreatic duct, the superior mesenteric artery and thesplenic artery. Arrows show the direction of flow and the territory tobe rinsed with detergent.

FIG. 12. Gross appearance of the human pancreas post-decellularization.In the case of human organs, the scaffold or acellular organ does notlook perfectly white or whitish.

FIG. 13. Staining of the head of the post-decellularization pancreaswith H&E, Masson Tric, Sirious Red and DAPI. All stains show the absenceof cells.

FIG. 14. Staining of the body of the post-decellularization pancreaswith H&E, Masson Tric, Sirious Red and DAPI. All stains show the absenceof cells.

FIG. 15. Staining of the tail of the post-decellularization pancreaswith H&E, Masson Tric, Sirious Red and DAPI. All stains show the absenceof cells.

FIG. 16. Comparison between the normal pancreas prior todecellularization and the pancreas after decellularization, revealingthe post-decellularization pancreas to be acellular.

FIG. 17. Various cell seeding methods for re-endothelialization ofacellular kidney scaffold. H&E stained images demonstrate thatcombination of static and ramping perfusion is the most effective methodto facilitate cell lining of the renal artery and vein (arrows) and theintermediate- and small-sized blood vessels (arrows) in the parenchymawithout cell clogging [(Small (cortex)]. The other three methodsresulted in low cell attachment on the different-sized blood vessels(arrowheads), with severe cell clogging (asterisk) in the cortex. Scalebars indicate 1 mm (artery and vein), 50 μm (magnified box ofartery/vein), and 200 μm (intermediate and small).

FIG. 18. Imaging of the re-endothelialized vascular tree. Lightmicroscopic images (a) show intact arterial branches in unseeded [EC(−)]and seeded kidney scaffold [EC(+)]. The green fluorescent image derivedfrom re-endothelialization was clearly visualized in [EC(+)] kidneyscaffolds, whereas there is no strong GFP signal in the [EC(−)]scaffolds (b). Fluorescent imaging of cortical surfaces (c) indicatesGFP-positive capillary-like structures in the [EC(+)] kidney scaffolds,indicating that viable MS1 cells are present in the cortical parenchyma.The SEM study indicates that the morphology of the endothelial cells ispreserved on the inner surface of artery (d). Scale bars: 200 μm (c) and50 μm (d).

FIG. 19. Imaging and quantification of platelet adhesion on thevasculatures of kidney scaffolds. Numerous integrin αIIb-positiveplatelets had adhered to the renal artery of [EC(−)] scaffolds (upperpanel in a) while the GFP positive MS1 coverage prevented plateletadhesion (lower panel in a), as indicated by arrows in the mergedimages. Quantitatively, the number of platelets adhered to the artery issignificantly lower in [EC(+)] than [EC(−)] scaffolds (*Student t-test,P<0.05, 3-4 sites per kidney, n=3 kidneys) (b). The fluorescentintensity from the integrin αIIb signals was determined to compare thelevels of platelet adhesion on intermediate- (c, d) and small-sized (e,f) blood vessels in the parenchyma. Similar to the results from therenal artery and vein, re-endothelialization dramatically reducedplatelet adhesion, and this was confirmed by quantitative analysis ofintermediate-size (d) (*Student t-test, P<0.05, three blood vessels perkidney, n=3 kidneys) and small size vessels (f) (*Student t-test,P<0.01, nine capillary structured vessels per kidney scaffold, n=3kidneys). Interestingly, three different sites of the cortex showed thistendency (f). Scale bars: 100 μm (a) and 50 μm (c,e). Data representmean±SD.

FIG. 20. In vivo implantation of MS1-seeded kidney scaffold into pigsfor 2 hours. (a) Before implantation, (b) surgical implantation, (c)ultrasound imaging at 1 hour after implantation, (d) the explantedscaffold from the implantation, and angiogram results of (e) before and(f) after injection of contrast agent, demonstrating severe blood clots(asterisks in g), as confirmed by histological examination (g, h). MS1detachment (arrows) and cell clogging (arrowheads) are observed in theimages of boxes in g and h. Renal artery (RA), renal vein (RV), inferiorvena cava (IVC) was shown in the panels. Scale bar: 100 μm (g), 200 μm(h), and 50 μm (box in g and h).

FIG. 21. Cell detachment assay under dynamic flow conditions todetermine the effect of antibody conjugation on improvement of celladherence. A microfluidic system (a) consisting of PDMS-based glasschamber was designed to examine the effect of antibody conjugation onthe cell detachment. The conjugated CD31 antibody was visualized as redfluorescent (c) while there is no significant brightness on theuntreated surfaces (b). Semi-quantification shows statistical differencebetween antibody-conjugated surfaces and untreated sample (*Studentt-test, P<0.05, n=3). Data represent mean±SD. In the cell detachmenttest (e, f), MS1 cells that pre-adhered on the Ab-conjugated surfaces[Ab(+)] maintained their cell number; however, cells on the untreatedsample [Ab(−)] went detached with the increase of flow rate and reachedto higher than 80% of originally pre-attached cells after flowing at 50ml/hr. Statistically significant differences were found at all points offlow rates (****, two-way ANOVA, P<0.00001, Post-hoc Bonferonni test,P<0.05 for all points, n=4). Scale bar: 200 μm (b,c). Data representmean±SEM.

FIG. 22. In vivo kidney implantation to determine the effect of CD31antibody conjugation of kidney scaffolds on the vascular maintenance.The re-endothelialized kidney scaffolds without [Ab(−)] and with [Ab(+)]antibody conjugation were implanted into pigs for 2 and 4 hours,respectively. The angiogram of the harvested kidney scaffolds showedmassive blood clots/occlusions and extravasation of host blood cells in[Ab(−)] (indicated by arrows), whereas a patent vascular tree wasobserved in [Ab(+)] (a). This functional outcome was confirmed byhistological examination, showing higher percentage of MS1endothelialization of [Ab(+)] than that of [Ab(−)] in the medulla site(intermediate-sized vessels, 3 sites per kidney, n=3 kidneys per group)and small-sized vessels in the cortex (9 sites per kidney, n=3 kidneysper group) (Student t-test,*P<0.05, **P=0.059) (b). The immunostainingusing platelet marker (═IIb integrin) showed numerous platelet adhesions(arrows) on the lumens of naked vasculatures of [Ab(−)], while lowadhesion was detected on the MS1-adhered intermediate and small bloodvessels of [Ab(+)] (c). The extent of platelet adhesion wassignificantly different with statistical analysis (*Student t-test,P<0.05) (d). Scale bar: 50 (intermediate) and 10 μm (small) (c). Datarepresent mean±SD.

FIG. 23. Schematic diagram of the overall re-endothelializationprocesses in the decellularized pig kidney scaffold and in vivoimplantation. Native kidney (a) was decellularized using 0.5% SDS andDNAase treatment to obtain a completely decellularized kidney scaffold(b). Following antibody conjugation on vasculatures of the kidneyscaffold (c), the antibody-conjugated scaffold was then seeded withendothelial cells (MS1) using a combination of the static (d) andramping perfusion methods (e), and then cultured in a bioreactor system.For in vivo function evaluation, the implantation of re-endothelializedkidney scaffold was performed by arterial [renal artery (RA) with aorta(A)] and venous [renal vein (RV) with inferior vena cava (IVC)]anastomosis into pigs (f). RA and RV are indicated by red and bluesuture material, respectively.

FIG. 24. Characterization of endothelial specificity by immunostainingusing endothelial cell markers to determine phenotypic maintenance ofMS1 cells. Endothelial cell markers such as CD31, vascular endothelialcadherin (VE-cad), and CD146 were used for the immunostaining. Nophenotypic change was detected. Scale bars: 50 μm.

FIG. 25. H&E histology of different-sized blood vessels in the renalparenchyma. The H&E staining demonstrates that the size of the bloodvessels decreased from segment 1 (closed renal artery and vein) tosegment 4 (mid-pole cortex) and most blood vessels were lined withseeded MS1 cells as shown in the magnified images. Particularly,small-sized vessels showed patency in segment 6 (superior pole) and 8(inferior pole) of the cortex. Scale bars: 50 μm (magnified boxes).

FIG. 26. In vitro blood perfusion to examine the effects ofre-endothelialization on the blood clogging and thrombosis. After bloodperfusion and rinsing with PBS, gross images of the kidney scaffold (a)showed large blood clot-like objects (arrows) in the unseeded scaffold[EC(−)], while no such finding was detected on the surface of [EC(+)]scaffolds. H&E staining confirmed that severe blood clotting occurred in[EC(−)] scaffolds (arrows in B), whereas the re-endothelializedscaffolds showed intact blood vessels (arrowheads) with only mild bloodcell clogging (arrows) in the parenchymal spaces. Scale bars: 1 cm (a),500 μm [EC(−)], and 100 μm [EC(+)] in (b).

FIG. 27. Whole human kidney de-cellularization and vascular corrosioncast. FIGS. 40A-D shows kidney macroscopic frontal view duringwhole-kidney detergent-based de-cellularization protocol. Representativetime lapse of kidney decellularization at T0 (A) and after 16, 32, and48 hours (T1, T2 and T3) (B, C and D, respectively). Peristalticperfusion through the renal artery and the ureter (12.5 ml/minute), withthe renal vein used as vascular outlet, permits homogeneous and completecellular compartment removal with a slow progressive change of the colorof the organ. FIGS. 40E-H shows a human renal vascular corrosion castobtained by injection of a specific resin via the renal artery (reddye), the renal vein (blue dye) and the ureter (yellow dye). Macroscopicanterior view of the endocast (E) shows the uniform distribution of theresin, obtained by a gentle continuous injection, inside the nativekidney vascular tree from the renal hilum to last vascular branching.Panel F shows the three-dimensional endocast of final renal vascularbranching system; the resin filled the finest branches of the vascularnetwork (arterial and venous). Panel G shows the complete preservationof the macrostructures of the organ (arterial, venous and ureteralcollecting system). The sample in question possesses a double-ureter andpelvicaliceal system that preserves its entire structure at alldetectable hierarchical levels. Even without cells present, the castingresin filled the whole scaffold in its upper and lower pole, producing afinal high-fidelity tridimensional reconstruction. Magnification of thelower pole of the endocast (H) shows preservation of the final vasculardivisions of the scaffold as observed from the native organ.

FIG. 28. Glomerular Scanning Electron Microscopy—SEM—morphometricanalysis. FIGS. 41A and 41C show representative SEM images of a singleextracted native glomerulus derived from the cortical portion of thevascular corrosion endocast of a native kidney, while FIGS. 41B and 41Dshow representative SEM images of a single extracted glomerulus derivedfrom the cortical portion of the vascular corrosion endocast of anacellular kidney (scalebar: 100 μm). In both series major glomerularcomponents (afferent artery and branched capillaries) are clearlyvisible. Background from the original snapshot (A and B) was removed toenhance and highlight effective glomerular morphological shape and tomore accurately extrapolate its morphometric measurements. Diameter wasmeasured twice, sagittally and transversally, (C and D) and then theaverage was calculated (green lines and circle). Afferent arteriolarwidth was acquired from three different transversal points (red lines)(C and D) and the final average assessed. Finally, six differenttransversal measurements were obtained (yellow lines) from singularcapillaries in several areas of the glomerulus (C and D).

FIG. 29. Morphometric analysis of native and de-cellularized glomeruli.The graphs present in FIG. 3 exemplify the mean measurements expressedin um and um³, of glomerular diameter (A), glomerular volume (B),afferent arteriolar diameter (C) and glomerular capillary with between80 glomeruli derived from native kidney cast samples and 80de-cellularized glomeruli derived from kidney cast samples. Whileglomerular diameter and volume significantly decrease in afterde-cellularization, no change was noticed in arteriolar and capillarydimensions before and after de-cellularization. ** p<0.01.

FIG. 30. Measurement of mean systolic (Max) and diastolic (Min) pressureresponsiveness. Panel A shows the measurement of mean systolic pressureresponsiveness of nine human kidneys (before and afterde-cellularization) machine-perfused at 90 mm Hg for 12 hours. Pointsrepresent the percent difference in the machine-set pressure and actualpressure. Native kidney, (blue line) and de-cellularized kidney, hrECMs(orange line) demonstrated an elastic response to machine set pressures(black line). The mean vascular elastic response in the native kidneywas 0.898% versus 1.76% after de-cellularization. Panel B shows themeasurement of mean diastolic pressure responsiveness of nine humankidneys (before and after de-cellularization) machine-perfused at 50 mmHg for 12 hours. Points represent the percent difference in themachine-set pressure and actual pressure. Native kidney (blue line) andde-cellularized kidney, hrECMs (orange line) demonstrated an elasticresponse to machine set pressures (black line). The mean vascularelastic response in the native kidney was 7.47% versus 2.48% afterde-cellularization. Presence of an elastic rebound in response to theapplied pulse wave confirmed that de-cellularization diminished but didnot compromise the elastic response of the vascular scaffold.

FIG. 31. GFs retention within hrECMs and ad hoc immunofluorescenceanalysis. Panel A shows the results of a semi-quantitative analysis ofthe protein expression profile of different GFs measured within thede-cellularized kidney (hrECMs) using a custom RayBio® Human GrowthFactors Antibody Array G series 1. Average fluorescent intensity wasobtained from duplicate signal intensities, adjusted to removebackground and normalized to a positive control to account fordifferences among sub-arrays. As a general trend, important moleculeslike TGFβ, IGFII, members of the TGF family and VEGF family are retainedwithin the decellularized matrix. Immunofluorescence for collagen V (B,20×), vascular endothelial growth factor receptor 2 (VEGFR2, C, 20×) andVe-Cadherin (D, 20×) demonstrate that the decellularized glomerulimaintain the presence of important fibrous protein (collagen V) as wellas VEGFR2 and Ve-Cadherin that are necessary for endothelial cellattachment and function.

FIG. 32. Schematic diagram of the re-endothelialization processes fordecellularized pig liver scaffolds. (A) Native liver harvested frompiglets was decellularized using 1% Triton X100 to obtain a completelydecellularized liver scaffold (B). Following antibody conjugation onvasculatures of the liver scaffold (C), the antibody-conjugated scaffoldwas seeded with endothelial cells (MS1) using a combination of static(D) and perfusion methods (E), and then matured in a bioreactor system.The engineered liver construct was implanted into pigs. PV, HA, SH-IVC,and IH-IVC indicates portal vein, hepatic artery, suprahepatic inferiorVena Cava, and intrahepatic inferior Vena Cava, respectively.

FIG. 33. Preparation of implantable porcine liver scaffold. (A) Grossimages of porcine liver decellularization, Ruler scale: 15 cm. (B)Evidence of decellularization of normal porcine liver (H&E staining andresidual DNA quantification). Acellular liver scaffolds have no cellularcomponents visible in the portal vein branch and central vein as well aswithin lobular structures. The quantification of residual DNA showedsignificant reduction in DNA as compared to that of native liver (n=3liver scaffolds per group, Student t-test, P<0.05). (C) Maintenance ofintact and functional vasculatures within the decellularized porcineliver scaffold by CT imaging and vascular casting followed by SEManalysis. CT images indicate intact vasculatures throughout entire lobesof the decellularized liver scaffold. SEM analyses of liver castsdemonstrate functional vasculatures. Note the sinusoid-like structures(arrows in the images of ×500). (D) Antibody conjugation ontovasculatures within the liver scaffold. CD31 antibody conjugation wasconfirmed by positive fluorescence derived from fluorescently labeledsecondary antibody. Quantitative results of antibody conjugation showedsignificant statistical difference in blood vessels of various sizes(Student t-test, P<0.05, n=3 liver scaffolds per group). Six large (>200mm) and three small (<200 mm) blood vessels per scaffold were selectedfor the quantification.

FIG. 34. Structural and functional re-endothelialization of acellularliver scaffolds. (A) Homogenous and well-oriented re-endothelializationof the portal vein (PV) of the decellularized liver scaffold, confirmedby GFP expression and SEM analysis on attached MS1 cells, respectively.(B) Efficient re-endothelialization at the portal triad and central vein(GFP/DAPI staining). (C) Functional re-endothelialization on thevasculature. Blood perfusion of liver scaffolds results in significantlevels of platelet adhesion (aIIb positive staining) on the vasculaturesof the unseeded scaffold (EC⁻) indicated by arrows (severe plateletadhesion), while endothelial cell attachment (arrowheads) wassignificantly reduced in the EC⁺ scaffolds. Quantitatively,re-endothelialization of liver scaffolds (EC

) significantly reduced platelet adhesion as compared with that ofscaffold only. The quantitative results showed significant statisticaldifference (Student t-test, P<0.05, n=3 liver scaffolds per group, fourareas from PV and CV per scaffold).

FIG. 35. In vivo implantation of engineered porcine liver constructsinto pigs. (A) The engineered liver construct was heterotopicallyimplanted into pigs (B) and showed good blood flow through the entireliver implant, as confirmed by ultrasound imaging at the anastomosissites of (C) renal artery (RA) and portal vein (PV), (D) infrahepatic(IH)-IVC and renal vein (RV), and each lobe [(E) LL and (F) RL lobe)].At 4 hours after implantation, the engineered liver implant demonstratedvascular patency, as evidenced by fluoroscopic angiography (GeI). LL andRL represents left and right lateral lobe, respectively.

FIG. 36. Improved vascular patency by re-endothelialization of the liverscaffold. At one day post-implantation, re-endothelialization ofvasculatures within the implant maintained vascular patency as comparedwith that of scaffold only, as confirmed by (A) ultrasound imaging aswell as (B) fluoroscopic angiography. (C) Results of H&E andimmunostaining for platelet adhesion demonstrate that MS1 attachment(arrowhead) on the vasculature significantly reduced platelet adhesiononto vessel walls of the re-endothelialized implants compared with thaton scaffold only (arrows). The quantitative results showed significantstatistical difference (Student t-test, P<0.05, n=3 liver scaffolds pergroup, four areas from PV and CV per scaffold).

FIG. 37. De-cellularization process, fluoroscopy and DNA quantitation.Panels A and B show the set-up of the core for organ decellularizationin a discarded human pancreas before and after the treatment. Red arrowsin the first image show the arterial, represented by superior mesentericartery (SMA) and splenic artery (SA) that share one of the two detergentinflows. Green arrow indicates the second inflow through the pancreaticduct (PD). During decellularization, the color of the organmacroscopically changed from a golden brown color to straw yellow butwithout reaching the whitish color described for porcine pancreasscaffolds (20). Fluoroscopy of hpaECMs performed through SMA (Panel C),SA (panel D) and PD (panel E, F, G). Contrast media flows within theframework of the innate vasculature of hpaECMs without extravasation.Panel H confirms satisfactory cell and DNA clearance. Statisticalanalysis included t-test of fresh versus acellular tissue; *** p<0,001.

FIG. 38. Basic histology of hpaECMs as compared to the native pancreas.H&E demonstrates complete loss of cellular elements with preservation ofthe intercellular framework. Without wishing to be bound by theory, thelarger empty spaces on the left aspect of the section may reflectpathological lesions, possibly representing focal fat necrosis. Massontrichrome shows the light green ECM framework. On the left aspect of theright panel there is evidence of scarring, indicated by the dense areastaining green. Alcian blue demonstrates preservation of the groundsubstance in the connective tissue framework, although the process ofdigestion involved in de-cellularization has altered the faint bluecolor of Alcian blue into a faint green color, as can be seen in theright panel. The van Gieson stain for elastic tissue demonstratespreservation of the elastic tissue of a couple of small structure ofunclear nature in the right panel. The picrosirius red demonstratesabundant collagen in the right panel, reflecting significant scarring inthe field photographed in contrast to the left panel.

FIG. 39. Immunohistochemistry. Aside of nuclear staining, presence ofpossible immunogens was ruled out by immunostaining for majorhistocompatibility complexes of class I and II (MHC I and II).Immunostaining for critical, ubiquitous components of pancreas ECMshowed that laminin, fibronectin, collagen type I and IV remainwell-preserved in hpaECMs (nuclei were counterstained with DAPI) anddisplays a similar distribution of these proteins in the fresh andde-cellularized pancreas bioscaffold sections.

FIG. 40. SEM of intact and acellular pancreas ECM. A. Low powermagnification of a cross section through the de-cellularized pancreasdemonstrating preservation of the ECM supporting the exocrine andendocrine elements as well as the vasculature. B. Three dimensional viewof the ECM framework of an islet of Langerhans in the de-cellularizedpancreas, identifiable by what is presumably its spherical microvascularframework. C. Cross-section through an arteriole with the lumendelimited by the internal elastic membrane and external to it, theindividual layers of basement membrane of dissolved smooth muscle fibersof the arteriole's media. D. A venule identifiable by the smaller numberof layers of smooth muscle cells (identified by the layers of basementmembrane) forming its wall and two small tributaries contributing to itsformation.

FIG. 41. CAM assay. CAM assay showed that, after seven days ofimplantation, the number of vessels converting towards thede-cellularized pancreas significantly increased in comparison to thesame samples at day 0, and the negative control.

FIG. 42. Seeding of islets onto de-cellularized scaffolds. (A)De-cellularized scaffolds prior to seeding. (B) Scaffold immediatelyafter seeding, black arrows indicate islets. (C) Dithizone-stainedislets after 4 days in culture on culture plates. (D) Dithizone-stainedislet located on periphery of scaffold, 4 days after seeding ontoscaffolds. The plot (E) illustrates dynamic perifusion results for humanislets after 4 days if culture. Here, both islets in standard cultureconditions as well as islets seeded statically on scaffoldingdemonstrated similar responses to changes in glucose where insulinsecretion rates increased during the 16.7 mM glucose perifusion, thendropped back down to basal levels during the second 3.3 mM glucoseperifusion period. Of 3 separate perifusion experiments the averagestimulation index for islets cultured on the scaffolds was 2.81±0.48which was not significantly different from islets cultured on standardplastic petri dishes, which had a stimulation index of 3.44±1.821(p=0.59, average±standard deviation, n=3).

FIG. 43. Endothelial cell seeding. Matrix was seeded with humanpancreatic endothelial cells and cultured for six days in a bioreactor,consisting of a closed circuit with one chamber for organ housing, areservoir for medium oxygenation and a peristaltic pump (Ismatec),connected by tubing (ID 1/16″, Pharmed BPT). Pancreatic tail wassurgically isolated in order to obtain a smaller volume to seed, keepingat the same time an inflow (SA—red connector) and an outflow (SplenicVein 9SV9 blue connector) (Panel A). Panel B indicates a schematicrepresentation of the perfusion circuit for seeded pancreatic scaffoldculture. Panel C shows a representative image of H&E stain showinglocalization of infused cells in vessels. Boxes indicate areas reportedwith high magnification in the panel below. Panel D illustratesrepresentative images of H&E (left), CD31 (middle) and Ki67 (right)matrix staining.

FIG. 44. Mechanical properties of hpaECMs. Graph on mechanicalproperties show similar patterns between native pancreata (non-treated)and hpaECM for tensile strain. Table below indicates data related tomodulus (expressed in MPa), tensile strain at break (%) and tensilestress at break (%).

DETAILED DESCRIPTION

The disclosure provides materials and methods for decellularizingtissues and organs ex vivo, producing organ and tissue scaffolds orframeworks composed of extracellular matrix containing intactvasculature suitable for recellularization. Thus, the disclosurecomprehends materials and methods for the decellularization of tissuesor organs, materials and methods for maintenance of the tissue or organscaffold, and materials and methods for the recellularization of suchtissue or organ scaffolds. The technology provides an opportunity toeliminate the cost, suffering, loss in quality of life and loss of lifeitself associated with the ever-increasing disparity between patients inneed of new tissues or organs and the supply thereof.

A pool of legally proper organs discarded for medical reasons may beused as a platform for renal bioengineering and regeneration research.The decellularization of porcine kidneys yields renal extracellularmatrix (ECM) scaffolds that maintain their basic components, supportcell growth and welfare in vitro and in vivo, and show an intactvasculature that, when such scaffolds are implanted in vivo, is able tosustain physiological blood pressure. Disclosed herein is a studyexpanding the strategy to discarded human kidneys in order to obtainhuman renal ECM scaffolds. The results show that the sodium dodecylsulfate-based decellularization protocol completely cleared the cellularcompartment in these kidneys, while the innate ECM framework retainedits architecture and biochemical properties. Samples of human renal ECMscaffolds stimulated angiogenesis in a chick chorioallantoic membraneassay. The innate vascular network in the human renal ECM scaffoldsretained its compliance. Collectively, these results indicate thatdiscarded human kidneys are a suitable source of renal scaffolds andtheir use for tissue engineering applications may be more clinicallyapplicable than kidneys derived from animals.

In attempts to bioengineer renal organ(oid)s for human transplantpurposes, whole, intact scaffolds produced from the extracellular matrix(ECM) of animal or human kidneys offer some promise. An advantage ofusing natural scaffolds from innate organs lies in the fact that suchbiomaterials are intrinsically functional when used as tissueengineering scaffolds, have their basic biochemistry (proteins andpolysaccharides) and architecture preserved, retain an intact and patentvasculature, and are able to drive differentiation of progenitor cellsinto organ-specific phenotypes.

After successful investigations in rodents [7-9, 13-17], the resultswere transferred to develop a more clinically relevant porcine model[25] because the evolutionary gap between humans and animal models isreduced with a pig model, although the use of non-human animal modelshas often prevented direct applicability of the knowledge toapplications in humans [26]. As noted herein, the United States alonediscards more than 2,600 kidneys annually from the total number ofkidneys procured from deceased donors. Disclosed herein are studiesassessing the suitability of human kidneys discarded fromtransplantation for use in renal bioengineering and organ(oid)regeneration. The main reasons for discard are severe anatomicalanomalies such as glomerulosclerosis, tubular atrophy, interstitialfibrosis, inflammation, cortical necrosis, and vascular changes, howeverother causes including prolonged cold ischemia, excessive warm ischemiaand poor renal function may intervene [27]. This number of organsaccounts for nearly 20% of the whole kidney pool procured for transplantpurposes, compared with about 12% in the late 1990s, and represent adramatic figure, at a time of severe organ shortage and urgent need forthe identification of new, valuable sources of organs. While the UnitedNetwork for Organ Sharing is currently devising strategies to minimizethe kidney discard rate, the technology disclosed herein salvages thosediscarded kidneys to produce ECM scaffolds for renal bioengineering andregeneration purposes.

The findings disclosed herein show that discarded human kidneys can besuccessfully decellularized to produce acellular renal ECM scaffolds.The acellular renal ECM scaffolds maintain their innate molecular andspatial framework, and are theoretically non-immunogenic because of theremoval of antigens that are responsible for the activation of theimmune rejection response. Thus, these renal scaffolds may represent anideal platform for investigations aimed at the manufacture of kidneysfor transplant purposes. Importantly, these scaffolds derive fromdiseased organs that were deemed unsuitable for clinical transplantationfor a variety of reasons, mainly severe biopsy-proven damage. In fact,the most frequent lesion we have noticed in this study was above 20%glomerulosclerosis, which is often associated with moderate to severevascular changes, interstitial fibrosis and tubular atrophy.

Studies in the porcine model have shown that the cellular compartment ofthe porcine kidneys can be completely cleared with SDS-based solutionsto ultimately generate acellular ECM scaffolds. These scaffolds retaintheir complex three-dimensional architecture and basic molecularcomponents. Their innate vascular tree remains intact at allhierarchical levels, including the tuft vessels and intact nephronframework, which can be easily identified using histology and electronicmicroscopy. The data also establish that these scaffolds can besuccessfully implanted in porcine recipients, using sequentialanastomosis of the renal vein and artery to the vena cava and abdominalaorta, respectively, and that the scaffold's vasculature sustains wellphysiological blood pressure.

The protocol applied in the studies disclosed herein for discarded humankidneys differs from the one used for pigs in that we perfuse detergentsnot only through the artery but also through the ureter retrograde intothe collecting system. We opted for this approach following initialunsuccessful attempts, while using only arterial perfusion, in which weobserved that the severe tissue damage appeared to impair thedecellularization process. It should be emphasized that when weperformed the visualization of the vascular network by injectingcontrast media within the renal artery, contrast cleared through therenal vein but no contrast leaked from the ureter. This indicates thatthe urinary tract compartment—which eventually drains into the ureter—issealed, i.e., totally separated, from the vascular network; the reasonfor which we elected to add retrograde ureteral rinsing and perfusion tomake sure that the detergent solution was delivered into all renalcompartments to allow successful decellularization.

The successful preparation of acellular human renal ECM scaffolds isemphasized by 1) maintenance of native renal architecture both and themacro and micro levels, including the vascular and nephron components,2) removal of important cell-associated immunogenic markers and 3)preservation of natural renal ECM proteins that have essential role intissue development and regeneration. In addition, the capability of thescaffolds to induce neoangiogenesis may be interpreted as evidence of exvivo—or, better, in ovo—regenerative capacity. Furthermore, fortransplantation applications, our findings indicate that: (a) thepreserved vasculature of acellular human renal ECM scaffolds is patentand resilient, as demonstrated by the fact that the vascular treemaintains the capability to modulate pressure following an increase ofthe inflow; (b) vessel resilience is determined in large part by theframework of the vasculature rather than its cellular compartment; (c)baseline pressure values are inferior to normal kidneys possibly due tothe lack of muscular layer within the vessel wall and/or the baselinediseased status of the vascular tree; d) the angiogenic capability ofthe ECM scaffold indicates that it can further support the vascularendothelium of the regenerated vascular network and thus improve themicrovasculature perfusion.

The results disclosed herein show that the sodium dodecyl sulfate-baseddecellularization protocol completely cleared the cellular compartmentin these kidneys, while the innate ECM framework retained itsarchitecture and biochemical properties. Samples of human renal ECMscaffolds stimulated angiogenesis in a chick chorioallantoic membraneassay. Importantly, the innate vascular network in the human renal ECMscaffolds retained its compliance. Collectively, these results indicatethat discarded human kidneys are a suitable source of renal scaffoldsand their use for tissue engineering applications is expected to be moreclinically applicable than kidneys derived from animals.

The strategy of using discarded organs introduces a new type ofbiomaterial, represented by natural scaffolds obtained from themanipulation of organs with pre-existing damage. It is expected thathuman renal ECM scaffolds from discarded organs will maintain theability to provide cell attachments in view of the successful seeding ofcells that should be followed by appropriate attachment, proliferationand function. As one of the cell's functions is ECM remodeling bycontinuous degradation followed by synthesis and deposition of new ECM,it is expected that the baseline fibrosis will be eventually reverted,thereby paving the way for successful regeneration of normal kidneytissue.

Facilitating the use of natural scaffolds to provide functional tissuesand organs through recellularization is the association of cell-bindingantibody products, e.g., an anti-CD31 antibody product, with thescaffolds, improving the ability of scaffolds to recellularize. Thus,disclosed herein are antibody products derived from an antibody thatspecifically binds to an endothelial cell or in induced pluripotent stemcell. An exemplary antibody product comprises each of the sixcomplementarity determining regions of the above-described antibody, inaddition to further sequences which provide a framework to support athree-dimensional conformation that retains the binding property of theantibody from which the antibody product is derived. In exemplaryembodiments, the antibody product comprises one or both of the lightchain variable region and/or the heavy chain variable region of anantibody binding to an endothelial cell or an iPS cell. When present,the light and heavy chain variable regions may be joined via a linker.Suitable linkers are known in the art and include, e.g., a linkercomprising a short amino acid sequence of about 5 to about 25 aminoacids, e.g., about 10 to about 20 amino acids.

In exemplary embodiments, the antibody product provided herein furthercomprises additional amino acid sequences. The binding agent may furthercomprise a constant region of a heavy chain and/or a constant region ofa light chain. Sequences for heavy and light chain constant regions arepublically available. For example, the National Center of BiotechnologyInformation (NCBI) nucleotide database provides a sequence of theconstant region of the IgG1 kappa light chain. See GenBank Accession No.DQ381549.1, incorporated herein by reference. Also, for example, theNCBI nucleotide database provides a sequence of the constant region ofthe Mus musculus IgG1. See GenBank Accession No. DQ381544.1.

The antibody product may be derived from an antibody of any type ofimmunoglobulin that is known in the art. For instance, the antibody canbe of any isotype, e.g., IgA, IgD, IgE, IgG, IgM. The antibody can bemonoclonal or polyclonal. The antibody can be a naturally occurringantibody, i.e., an antibody isolated and/or purified from a mammal,e.g., mouse, rabbit, goat, horse, chicken, hamster, human, and the like.In this regard, the antibody may be considered to be a mammalianantibody, e.g., a mouse antibody, rabbit antibody, goat antibody, horseantibody, chicken antibody, hamster antibody, human antibody, and thelike. The term “isolated” as used herein means having been removed fromits natural environment. The term “purified,” as used herein relates tothe isolation of a molecule or compound in a form that is substantiallyfree of contaminants normally associated with the molecule or compoundin a native or natural environment and means having been increased inpurity as a result of being separated from other components of theoriginal composition. It is recognized that “purity” is a relative term,and not to be necessarily construed as absolute purity or absoluteenrichment or absolute selection. In some aspects, the purity is atleast or about 50%, is at least or about 60%, at least or about 70%, atleast or about 80%, or at least or about 90% (e.g., at least or about91%, at least or about 92%, at least or about 93%, at least or about94%, at least or about 95%, at least or about 96%, at least or about97%, at least or about 98%, at least or about 99% or is approximately100%. In exemplary aspects, the antibody comprises a constant region ofan IgG, such as the constant region of an IgG₁. The antibody maycomprise the constant region of an IgG kappa light chain. In someembodiments, the antibody comprises a constant region of a Mus musculusIgG₁.

The antibody products of the disclosure can have any level of affinityor avidity for an endothelial cell or an iPS cell. In certainembodiments in which the antibody product comprises two or more distinctantigen binding regions or fragments, the antibody product is consideredbispecific, trispecific, or multi-specific, or bivalent, trivalent, ormultivalent, depending on the number of distinct epitopes that arerecognized and bound by the antibody product. The antibody product inexemplary aspects is considered to be a blocking antibody orneutralizing antibody.

In exemplary embodiments, the antibody product is a geneticallyengineered antibody, e.g., a single chain antibody, a humanizedantibody, a chimeric antibody, a CDR-grafted antibody, an antibody thatincludes portions of CDR sequences specific for a cell-surface marker ofan endothelial cell or an iPS cell, a humaneered or humanized antibody,a bispecific antibody, a trispecific antibody, and the like, as definedin greater detail herein. Genetic engineering techniques also providethe ability to make fully human antibodies in a non-human.

In some aspects, the antibody product is a chimeric antibody. The term“chimeric antibody” is used herein to refer to an antibody containingconstant domains from one species and the variable domains from asecond, or more generally, containing stretches of amino acid sequencefrom at least two species.

In some aspects, the antibody is a humanized antibody. The term“humanized” when used in relation to antibodies is used to refer toantibodies having at least CDR regions from a nonhuman source that areengineered to have a structure and immunological function more similarto true human antibodies than the original source antibodies. Forexample, humanizing can involve grafting CDR from a non-human antibody,such as a mouse antibody, into a human antibody. Humanizing also caninvolve select amino acid substitutions to make a non-human sequencelook more like a human sequence, as would be known in the art.

Use of the terms “chimeric or humanized” herein is not meant to bemutually exclusive; rather, is meant to encompass chimeric antibodyproducts, humanized antibody products, and chimeric antibody productsthat have been further humanized. Except where context otherwiseindicates, statements about (properties of, uses of, testing, and so on)chimeric antibody products apply to humanized antibody products, andstatements about humanized antibody products pertain also to chimericantibody products.

In some aspects of the disclosure, the antibody product is an antigenbinding fragment of an antibody that specifically binds to anendothelial cell or iPS cell in accordance with the disclosure. Theantigen binding fragment (also referred to herein as “antigen bindingportion”) may be an antigen binding fragment of any of the antibodiesdescribed herein. The antigen binding fragment can be any part of anantibody that has at least one antigen binding site including, but notlimited to, Fab, F(ab′)₂, dsFv, sFv, diabodies, triabodies, bis-scFvs,fragments expressed by a Fab expression library, domain antibodies, VhHdomains, V-NAR domains, VH domains, VL domains, and the like. Antibodyfragments of the disclosure, however, are not limited to these exemplarytypes of antibody fragments.

In exemplary aspects, the antibody product comprises a leader sequence.In exemplary aspects, the antigen binding fragment comprises an Ig kappaleader sequence. Suitable leader sequences are known in the art.

In exemplary aspects, an antibody product of the disclosure comprisesone more tag sequences. Tag sequences may assist in the production andcharacterization of the manufactured antigen binding fragment. Inexemplary aspects, the antigen binding fragment comprises one or moretag sequences C-terminal to the binding domains of the antibody product.Suitable tag sequences are known in the art and include, but are notlimited to, Myc tags, His tags, and the like.

In exemplary aspects, an antibody product of the disclosures comprises,from the N- to the C-terminus, a leader sequence, a heavy chain variableregion, a linker sequence, a light chain variable region, a Myc tag, anda His tag.

In exemplary aspects, the antibody product is a domain antibody. Adomain antibody comprises a functional binding unit of an antibody, andcan correspond to the variable regions of either the heavy (V_(H)) orlight (V_(L)) chains of antibodies. A domain antibody can have amolecular weight of approximately 13 kDa, or approximately one-tenth theweight of a full antibody. Domain antibodies may be derived from fullantibodies. The antibody products in some embodiments are monomeric orpolymeric, bispecific or trispecific, and bivalent or trivalent.

Antibody products that contain the antigen binding, or idiotope, of anantibody molecule share a common idiotype and are contemplated by thedisclosure. Such antibody products may be generated by techniques knownin the art and include, but are not limited to, the F(ab′)₂ fragmentwhich may be produced by pepsin digestion of an antibody; the Fab′fragments which may be generated by reducing the disulfide bridges of aF(ab′)₂ antibody, and the two Fab′ fragments which may be generated bytreating an antibody with papain and a reducing agent.

In exemplary aspects, the antibody product provided herein is asingle-chain variable region fragment (scFv) antibody fragment. An scFvmay consist of a truncated Fab fragment comprising the variable (V)domain of an antibody heavy chain linked to a V domain of an antibodylight chain via a synthetic peptide, and it can be generated usingroutine recombinant DNA technology techniques (see, e.g., Janeway etal., Immunobiology, 2^(nd) Edition, Garland Publishing, New York,(1996)). Similarly, disulfide-stabilized variable region fragments(dsFv) can be prepared by recombinant DNA technology (see, e.g., Reiteret al., Protein Engineering, 7, 697-704 (1994)).

Recombinant antibody products, e.g., scFvs of the disclosure, can alsobe engineered to assemble into stable multimeric oligomers of highbinding avidity and specificity to different target antigens onendothelial and iPS cells. Such diabodies (dimers), triabodies (trimers)or tetrabodies (tetramers) are well known in the art. See e.g., Kortt etal., Biomol Eng. 2001 18:95-108, (2001) and Todorovska et al., J ImmunolMethods. 248:47-66, (2001).

In exemplary embodiments, the antibody product is a bispecific antibody(bscAb). Bispecific antibodies are molecules comprising two single-chainFv fragments joined via a glycine-serine linker using recombinantmethods. The V light-chain (V_(L)) and V heavy-chain (V_(H)) domains oftwo antibodies of interest in exemplary embodiments are isolated usingstandard PCR methods. The V_(L) and V_(H) cDNAs obtained from eachhybridoma are then joined to form a single-chain fragment in a two-stepfusion PCR. Bispecific fusion proteins are prepared in a similar manner.Bispecific single-chain antibodies and bispecific fusion proteins areantibody substances included within the scope of the present invention.Exemplary bispecific antibodies are taught in U.S. Patent ApplicationPublication No. 2005-0282233A1 and International Patent ApplicationPublication No. WO 2005/087812, both applications of which areincorporated herein by reference in their entireties.

In exemplary embodiments, the antibody product is a bispecific T-cellengaging antibody (BiTE) containing two scFvs produced as a singlepolypeptide chain. Methods of making and using BiTE antibodies aredescribed in the art. See, e.g., Cioffi et al., Clin Cancer Res 18: 465,Brischwein et al., Mol Immunol 43:1129-43 (2006); Amann M et al., CancerRes 68:143-51 (2008); Schlereth et al., Cancer Res 65: 2882-2889 (2005);and Schlereth et al., Cancer Immunol Immunother 55:785-796 (2006).

The antibody product may be a dual affinity re-targeting antibody(DART). DARTs are produced as separate polypeptides joined by astabilizing interchain disulfide bond. In exemplary embodiments, theantibody product is a DART comprising an scFv. Methods of making andusing DART antibodies are described in the art. See, e.g., Rossi et al.,MAbs 6: 381-91 (2014); Fournier and Schirrmacher, BioDrugs 27:35-53(2013); Johnson et al., J Mol Biol 399:436-449 (2010); Brien et al., JVirol 87: 7747-7753 (2013); and Moore et al., Blood 117:4542 (2011).

In exemplary embodiments, the antibody product is a tetravalent tandemdiabody (TandAbs) in which an antibody fragment is produced as anon-covalent homodimer folded in a head-to-tail arrangement. TandAbs areknown in the art. See, e.g., McAleese et al., Future Oncol 8: 687-695(2012); Portner et al., Cancer Immunol Immunother 61:1869-1875 (2012);and Reusch et al., MAbs 6:728 (2014).

The BiTE, DART, or TandAbs of the disclosure comprise the CDRs of anantibody that binds an endothelial or iPS cell.

Suitable methods of making antibodies and/or antibody product are knownin the art. For instance, standard hybridoma methods are described in,e.g., Harlow and Lane (eds.), Antibodies: A Laboratory Manual, CSH Press(1988), and C A. Janeway et al. (eds.), Immunobiology, 5^(th) Ed.,Garland Publishing, New York, N.Y. (2001)).

Monoclonal antibodies as source material for the antibody products ofthe disclosure may be prepared using any technique that provides for theproduction of antibody molecules by continuous cell lines in culture.These include, but are not limited to, the hybridoma techniqueoriginally described by Koehler and Milstein (Nature 256: 495-497,1975), the human B-cell hybridoma technique (Kosbor et al., ImmunolToday 4:72, 1983; Cote et al., Proc Natl Acad Sci 80: 2026-2030, 1983)and the EBV-hybridoma technique (Cole et al., Monoclonal Antibodies andCancer Therapy, Alan R Liss Inc, New York N.Y., pp 77-96, (1985).

Briefly, a polyclonal antibody is prepared by immunizing an animal withan immunogen comprising an antigenic polypeptide and collecting antiserafrom that immunized animal. A wide range of animal species can be usedfor the production of antisera. In some aspects, an animal used forproduction of anti-antisera is a non-human animal including rabbits,mice, rats, hamsters, goat, sheep, pigs or horses. Because of therelatively large blood volume of rabbits, a rabbit, in some exemplaryaspects, is a preferred choice for production of polyclonal antibodies.In an exemplary method for generating a polyclonal antiseraimmunoreactive with the chosen marker of an endothelial and/or iPS cell,50 μg of antigenic marker is emulsified in Freund's Complete Adjuvantfor immunization of rabbits. At intervals of, for example, 21 days, 50μg of epitope are emulsified in Freund's Incomplete Adjuvant for boosts.Polyclonal antisera may be obtained, after allowing time for antibodygeneration, simply by bleeding the animal and preparing serum samplesfrom the whole blood.

Briefly, in exemplary embodiments, to generate monoclonal antibodies, amouse is injected periodically with the antigenic marker against whichthe antibody is to be raised (e.g., 10-20 μg of the marker emulsified inFreund's Complete Adjuvant). The mouse is given a final pre-fusion boostof the antigenic marker, and four days later the mouse is sacrificed andits spleen removed. The spleen is placed in 10 ml serum-free RPMI 1640,and a single-cell suspension is formed by grinding the spleen betweenthe frosted ends of two glass microscope slides submerged in serum-freeRPMI 1640, supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate, 100units/ml penicillin, and 100 μg/ml streptomycin (RPMI) (Gibco, Canada).The cell suspension is filtered through sterile 70-mesh Nitex cellstrainer (Becton Dickinson, Parsippany, N.J.), and is washed twice bycentrifuging at 200 g for 5 minutes and resuspending the pellet in 20 mlserum-free RPMI. Splenocytes taken from three naive Balb/c mice areprepared in a similar manner and used as a control. NS-1 myeloma cells,kept in log phase in RPMI with 11% fetal bovine serum (FBS) (HycloneLaboratories, Inc., Logan, Utah) for three days prior to fusion, arecentrifuged at 200 g for 5 minutes, and the pellet is washed twice.

Spleen cells (1×10⁸) are combined with 2.0×10⁷ NS-1 cells andcentrifuged, and the supernatant is aspirated. The cell pellet isdislodged by tapping the tube, and 1 ml of 37° C. PEG 1500 (50% in 75 mMHepes, pH 8.0) (Boehringer Mannheim) is added with stirring over thecourse of 1 minute, followed by the addition of 7 ml of serum-free RPMIover 7 minutes. An additional 8 ml RPMI is added and the cells arecentrifuged at 200 g for 10 minutes. After discarding the supernatant,the pellet is resuspended in 200 ml RPMI containing 15% FBS, 100 μMsodium hypoxanthine, 0.4 μM aminopterin, 16 μM thymidine (HAT) (Gibco),25 units/ml IL-6 (Boehringer Mannheim) and 1.5×10⁶ splenocytes/ml andplated into 10 Corning flat-bottom 96-well tissue culture plates(Corning, Corning N.Y.).

On days 2, 4, and 6, after the fusion, 100 μl of medium is removed fromthe wells of the fusion plates and replaced with fresh medium. On day 8,the fusion is screened by ELISA, testing for the presence of mouse IgGbinding to the marker.

Selected fusion wells are cloned twice by dilution into 96-well platesand visual scoring of the number of colonies/well after 5 days. Themonoclonal antibodies produced by hybridomas are isotyped using theIsostrip system (Boehringer Mannheim, Indianapolis, Ind.).

When the hybridoma technique is employed, myeloma cell lines may beused. Such cell lines suited for use in hybridoma-producing fusionprocedures preferably are non-antibody-producing, have high fusionefficiency, and enzyme deficiencies that render them incapable ofgrowing in certain selective media that support the growth of only thedesired fused cells (hybridomas). For example, where the immunizedanimal is a mouse, one may use P3-X63/Ag8, P3-X63-Ag8.653, NS1/1.Ag 4 1,Sp210-Ag14, FO, NSO/U, MPC-11, MPC11-X45-GTG 1.7 and S194/15XX0 Bul; forrats, one may use R210.RCY3, Y3-Ag 1.2.3, IR983F and 4B210; and U-266,GM1500-GRG2, LICR-LON-HMy2 and UC729-6 are all useful in connection withcell fusions. It should be noted that the hybridomas and cell linesproduced by such techniques for producing the monoclonal antibodies arecontemplated to be compositions of the disclosure.

Depending on the host species, various adjuvants may be used to increasean immunological response. Such adjuvants include, but are not limitedto, Freund's, mineral gels such as aluminum hydroxide, and surfaceactive substances such as lysolecithin, pluronic polyols, polyanions,peptides, oil emulsions, keyhole limpet hemocyanin, and dinitrophenol.BCG (bacilli Calmette-Guerin) and Corynebacterium parvum are potentiallyuseful human adjuvants.

Alternatively, other methods, such as EBV-hybridoma methods (Haskard andArcher, J. Immunol. Methods, 74(2), 361-67 (1984),and Roder et al.₅Methods Enzymol., 121, 140-67 (1986)), and bacteriophage vectorexpression systems (see, e.g., Huse et al., Science, 246, 1275-81(1989)) that are known in the art may be used. Further, methods ofproducing antibodies in non-human animals are described in, e.g., U.S.Pat. Nos. 5,545,806, 5,569,825, and 5,714,352, and U.S. PatentApplication Publication No. 2002/0197266 A1.

Antibodies may also be produced by inducing in vivo production in thelymphocyte population or by screening recombinant immunoglobulinlibraries or panels of highly specific binding reagents as disclosed inOrlandi et al. (Proc. Natl. Acad. Sci. 86: 3833-3837; 1989), and Winterand Milstein (Nature 349: 293-299, 1991).

Furthermore, phage display can be used to generate an antibody of thedisclosure. In this regard, phage libraries encoding antigen-bindingvariable (V) domains of antibodies can be generated using standardmolecular biology and recombinant DNA techniques (see, e.g., Sambrook etal. (eds.), Molecular Cloning, A Laboratory Manual, 3^(rd) Edition, ColdSpring Harbor Laboratory Press, New York (2001)). Phage encoding avariable region with the desired specificity are selected for specificbinding to the desired antigen, and a complete or partial antibody isreconstituted comprising the selected variable domain. Nucleic acidsequences encoding the reconstituted antibody are introduced into asuitable cell line, such as a myeloma cell used for hybridomaproduction, such that antibodies having the characteristics ofmonoclonal antibodies are secreted by the cell (see, e.g., Janeway etal., supra, Huse et al., supra, and U.S. Pat. No. 6,265,150). Relatedmethods also are described in U.S. Pat. Nos. 5,403,484; 5,571,698;5,837,500; and 5,702,892. The techniques described in U.S. Pat. Nos.5,780,279; 5,821,047; 5,824,520; 5,855,885; 5,858,657; 5,871,907;5,969,108; 6,057,098; and 6,225,447, are also contemplated as useful inpreparing antibodies according to the disclosure.

Antibodies can be produced by transgenic mice that are transgenic forspecific heavy and light chain immunoglobulin genes. Such methods areknown in the art and described in, for example U.S. Pat. Nos. 5,545,806and 5,569,825, and Janeway et al., supra.

Methods for generating humanized antibodies are well known in the artand are described in detail in, for example, Janeway et al., supra, U.S.Pat. Nos. 5,225,539; 5,585,089; and 5,693,761; European Patent No.0239400 B1; and United Kingdom Patent No. 2188638. Humanized antibodiescan also be generated using the antibody resurfacing technologydescribed in U.S. Pat. No. 5,639,641 and Pedersen et al., J. Mol. Biol.,235:959-973 (1994).

Techniques developed for the production of “chimeric antibodies,” thesplicing of mouse antibody genes to human antibody genes to obtain amolecule with appropriate antigen specificity and biological activity,can be used (Morrison et al., Proc. Natl. Acad. Sci. 81: 6851-6855,1984; Neuberger et al., Nature 312: 604-608, 1984; and Takeda et al.,Nature 314: 452-454; 1985). Alternatively, techniques described for theproduction of single-chain antibodies (U.S. Pat. No. 4,946,778) can beadapted to produce endothelial or iPS cell-specific single chainantibodies.

A preferred chimeric or humanized antibody has a human constant region,while the variable region, or at least a CDR, of the antibody is derivedfrom a non-human species. Methods for humanizing non-human antibodiesare well known in the art. (see U.S. Pat. Nos. 5,585,089, and5,693,762). Generally, a humanized antibody has one or more amino acidresidues introduced into a CDR region and/or into its framework regionfrom a source which is non-human. Humanization can be performed, forexample, using methods described in Jones et al. (Nature 321: 522-525,1986), Riechmann et al., (Nature, 332: 323-327, 1988) and Verhoeyen etal. (Science 239:1534-1536, 1988), by substituting at least a portion ofa rodent complementarity-determining region (CDR) for the correspondingregion of a human antibody. Numerous techniques for preparing engineeredantibodies are described, e.g., in Owens and Young, J. Immunol. Meth.,168:149-165 (1994). Further changes can then be introduced into theantibody framework to modulate affinity or immunogenicity.

Consistent with the foregoing description, compositions comprising CDRsmay be generated using, at least in part, techniques known in the art toisolate CDRs. Complementarity-determining regions are characterized bysix polypeptide loops, three loops for each of the heavy or light chainvariable regions. The amino acid position in a CDR is defined by Kabatet al., “Sequences of Proteins of Immunological Interest,” U.S.Department of Health and Human Services, (1983), which is incorporatedherein by reference. For example, hypervariable regions of humanantibodies are roughly defined to be found at residues 28 to 35, from49-59 and from residues 92-103 of the heavy and light chain variableregions [Janeway et al., supra]. The murine CDRs also are found atapproximately these positions in the heavy and light chain variableregions of antibodies. It is understood in the art that CDR regions maybe found within several amino acids of the approximated amino acidpositions set forth above. An immunoglobulin variable region alsoconsists of four “framework” regions surrounding the CDRs (FR1-4). Thesequences of the framework regions of different light or heavy chainsare highly conserved within a species, and are also conserved betweenhuman and murine sequences.

Compositions comprising one, two, and/or three CDRs of a heavy chainvariable region or a light chain variable region of a monoclonalantibody are generated. Polypeptide compositions comprising one, two,three, four, five and/or six complementarity-determining regions of anantibody are contemplated. Using the conserved framework sequencessurrounding the CDRs, PCR primers complementary to these consensusframework sequences are generated to amplify the CDR sequence locatedbetween the primer regions. Techniques for cloning and expressingnucleotide and polypeptide sequences are well-established in the art[see e.g., Sambrook et al., Molecular Cloning: A Laboratory Manual,2^(nd) Edition, Cold Spring Harbor, N.Y. (1989)]. The amplified CDRsequences are ligated into an appropriate plasmid. The plasmidcomprising one, two, three, four, five and/or six cloned CDRs optionallycontains additional polypeptide encoding regions linked to the CDR.

It is contemplated that modified polypeptide compositions comprisingone, two, three, four, five, or six CDRs of a heavy or light chain of animmunoglobulin are generated, wherein a CDR is altered to provideincreased specificity or affinity or avidity to the target IL13Rα2.Sites at locations in the CDRs are typically modified in series, e.g.,by substituting first with conservative choices (e.g., hydrophobic aminoacid substituted for a non-identical hydrophobic amino acid) and thenwith more dissimilar choices (e.g., hydrophobic amino acid substitutedfor a charged amino acid), and then deletions or insertions may be madeat the target site.

Framework regions (FR) of a murine antibody are humanized bysubstituting compatible human framework regions chosen from a largedatabase of human antibody variable sequences, including over twelvehundred human V_(H) sequences and over one thousand V_(L) sequences. Thedatabase of antibody sequences used for comparison is downloaded fromAndrew C. R. Martin's KabatMan web page(http://www.rubic.rdg.ac.uk/abs/). The Kabat method for identifying CDRsprovides a means for delineating the approximate CDR and frameworkregions of any human antibody and comparing the sequence of a murineantibody for similarity to determine the CDRs and FRs. Best-matchedhuman V_(H) and V_(L) sequences are chosen on the basis of high overallframework matching, similar CDR length, and minimal mismatching ofcanonical and V_(H)/V_(L) contact residues. Human framework regions mostsimilar to the murine sequence are inserted between the murine CDRs.Alternatively, the murine framework region may be modified by makingamino acid substitutions of all or part of the native framework regionthat more closely resembles a framework region of a human antibody.

“Conservative” amino acid substitutions are made on the basis ofsimilarity in polarity, charge, solubility, hydrophobicity,hydrophilicity, and/or the amphipathic nature of the residues involved.For example, nonpolar (hydrophobic) amino acids include alanine (Ala,A), leucine (Leu, L), isoleucine (Ile, I), valine (Val, V), proline(Pro, P), phenylalanine (Phe, F), tryptophan (Trp, W), and methionine(Met, M); polar neutral amino acids include glycine (Gly, G), serine(Ser, S), threonine (Thr, T), cysteine (Cys, C), tyrosine (Tyr, Y),asparagine (Asn, N), and glutamine (Gln, Q); positively charged (basic)amino acids include arginine (Arg, R), lysine (Lys, K), and histidine(His, H); and negatively charged (acidic) amino acids include asparticacid (Asp, D) and glutamic acid (Glu, E). “Insertions” or “deletions”are preferably in the range of about 1 to 20 amino acids, morepreferably 1 to 10 amino acids. The variation may be introduced bysystematically making substitutions of amino acids in a polypeptidemolecule using recombinant DNA techniques and assaying the resultingrecombinant variants for activity. Nucleic acid alterations can be madeat sites that differ in the nucleic acids from different species(variable positions) or in highly conserved regions (constant regions).

Additionally, another useful technique for generating antibodies for usein the methods of the disclosure may be one which uses a rationaldesign-type approach. The goal of rational design is to producestructural analogs of biologically active polypeptides or compounds withwhich they interact (agonists, antagonists, inhibitors, peptidomimetics,binding partners, and the like). In this case, the active polypeptidesare antibody products specifically binding an endothelial or iPS cell.By creating analogs of such polypeptides, it is possible to fashionadditional antibodies that are more immunoreactive than the native ornatural molecule. In one approach, one would generate athree-dimensional structure for the antibodies or an epitope bindingfragment thereof. This could be accomplished by x-ray crystallography,computer modeling or by a combination of both approaches. An alternativeapproach, “alanine scan,” involves the random replacement of residuesthroughout a molecule with alanine, and the resulting effect on functionis determined.

It also is possible to solve the crystal structure of the specificantibodies. In principle, this approach yields a pharmacore upon whichsubsequent drug design can be based. It is possible to bypass proteincrystallography altogether by generating anti-idiotypic antibodies to afunctional, pharmacologically active antibody. As a minor image of amirror image, the binding site of anti-idiotype antibody is expected tobe an analog of the original antigen. The anti-idiotype antibody is thenbe used to identify and isolate additional antibodies from banks ofchemically- or biologically-produced peptides.

Chemically synthesized bispecific antibodies may be prepared bychemically cross-linking heterologous Fab or F(ab′)₂ fragments by meansof chemicals such as heterobifunctional reagentsuccinimidyl-3-(2-pyridyldithiol)-propionate (SPDP, Pierce Chemicals,Rockford, Ill.). The Fab and F(ab′)₂ fragments can be obtained fromintact antibody by digesting it with papain or pepsin, respectively(Karpovsky et al., J. Exp. Med. 160:1686-701, 1984; Titus et al., J.Immunol., 138:4018-22, 1987).

Methods of testing antibodies for the ability to bind to endothelial oriPS markers, regardless of how the antibodies are produced, are known inthe art and include any antibody-antigen binding assay such as, forexample, radioimmunoassay (RIA), ELISA, Western blot,immunoprecipitation, and competitive inhibition assays (see, e.g.,Janeway et al., infra, and U.S. Patent Application Publication No.2002/0197266 A1).

Recent advances in the field of combinatorial sciences have identifiedaptamers, i.e., short polymer sequences (e.g., oligonucleic acid orpeptide molecules) with high affinity and specificity to a given target.For example, SELEX technology has been used to identify DNA and RNAaptamers with binding properties that rival mammalian antibodies. Thefield of immunology has generated and isolated antibodies or antibodyfragments which bind to a myriad of compounds, and phage display hasbeen utilized to discover new peptide sequences with very favorablebinding properties. Based on the success of these molecular evolutiontechniques, it is certain that molecules can be created which bind toany target molecule. A loop structure is often involved in providing thedesired binding attributes as in the case of aptamers, which oftenutilize hairpin loops created from short regions without complementarybase pairing, naturally derived antibodies that utilize combinatorialarrangement of looped hyper-variable regions and new phage-displaylibraries utilizing cyclic peptides that have shown improved resultswhen compared to linear peptide phage display results. Thus, sufficientevidence has been generated to indicate that high-affinity ligands canbe created and identified by combinatorial molecular evolutiontechniques, and molecular evolution techniques can be used to isolateantibody products specific for the endothelial and/or iPS cellsdisclosed herein. For more on aptamers, see generally, Gold, L., Singer,B., He, Y. Y., Brody. E., “Aptamers As Therapeutic And DiagnosticAgents,” J. Biotechnol. 74:5-13 (2000). Relevant techniques forgenerating aptamers are found in U.S. Pat. No. 6,699,843, which isincorporated herein by reference in its entirety.

In some embodiments, the aptamer is generated by preparing a library ofnucleic acids; contacting the library of nucleic acids with anendothelial or iPS cell marker, wherein nucleic acids having greaterbinding affinity for the marker (relative to other library nucleicacids) are selected and amplified to yield a mixture of nucleic acidsenriched for nucleic acids with relatively higher affinity andspecificity for binding to the marker. The processes may be repeated,and the selected nucleic acids mutated and rescreened, whereby a markeraptamer is identified. Nucleic acids may be screened to select formolecules that bind to more than target. Binding more than one targetcan refer to binding more than one simultaneously or competitively. Insome embodiments, an antibody product comprises at least one aptamer,wherein a first binding unit binds a first epitope of an endothelial oriPS cell marker and a second binding unit binds a second epitope of themarker.

With regard to the antibody products of the compositions of thedisclosure, attachment of endothelial and/or iPS cells to an organ ortissue scaffold is enhanced by immobilizing an antibody product to ascaffold and allowing the antibody product to facilitate the associationof endothelial and/or iPS cells with the scaffold. As used herein, theterm “facilitate” or “enhance” does not necessarily mean a 100% increasein the rate of association, the strength of association or the durationof association. Rather, there are varying degrees of facilitation orenhancement that one of ordinary skill in the art would recognize ashaving a potential benefit or therapeutic effect. Methods of measuringenhanced or facilitated association or attachment of endothelial or iPScells to tissue or organ scaffolds are known in the art.

The antibody products according to the disclosure specifically bind toat least one endothelial cell or iPS cell marker. Endothelial cellmarkers contemplated by the disclosure include CD31/PECAM-1, ACE/CD143,MCAM/CD146, C1q R1/CD93, Nectin-2/CD112, VE-Cadherin, PD-ECGF/ThymidinePhosphorylase, CC Chemokine Receptor D6, Podocalyxin, Podoplanin, CD34,S1P1/EDG-1, CD36/SR-B3, S1P2/EDG-5, CD151, S1P3/EDG-3, CD160,S1P4/EDG-6, CD300LG/Nepmucin, S1P5/EDG-8, CL-K1/COLEC11,E-Selectin/CD62E, CL-P1/COLEC12, E-Selectin (CD62E)/P-Selectin (CD62P),Coagulation Factor III/Tissue Factor, P-Selectin/CD62P, DC-SIGNR/CD299,SLAM/CD150, DCBLD2/ESDN, Stabilin-1, EMMPRIN/CD147, Stabilin-2,Endoglin/CD105, TEM7/PLXDC1, Endomucin, TEM8/ANTXR1, Endosialin/CD248,Thrombomodulin/BDCA-3, EPCR, THSD1, Erythropoietin R, Tie-2, ESAM, TNFRI/TNFRSF1A, FABP5/E-FABP, TNF RII/TNFRSF1B, FABP6, TRA-1-85/CD147,ICAM-1/CD54, TRAIL R1/TNFRSF10A, ICAM-2/CD102, TRAIL R2/TNFRSF10B, IL-1RI, VCAM-1/CD106, IL-13 R alpha 1, VE-Statin, Integrin alpha 4/CD49d,VEGF R1/Flt-1, Integrin alpha 4 beta 1, VEGF R2/KDR/Flk-1, Integrinalpha 4 beta 7/LPAM-1, VEGF R3/Flt-4, Integrin beta 2/CD18, VG5Q, KLF4,vWF-A2, or LYVE-1. An exemplary endothelial cell marker is CD31/PECAM-1.

Induced pluripotent stem (iPS) cell markers contemplated by thedisclosure include 5T4, Lefty, ABCG2, Lefty-1, Activin RIB/ALK-4,Lefty-A, Activin RIIB, LIN-28A, Alkaline Phosphatase/ALPL, LIN-28B,E-Cadherin, LIN-41, Cbx2, c-Maf, CD9, c-Myc, CD30/TNFRSF8, Nanog,CD117/c-kit, Oct-3/4, CDX2, Oct-4A, CHD1, Podocalyxin, Cripto,Rex-1/ZFP42, DNMT3B, Smad2, DPPA2, Smad2/3, DPPA4, SOX2, DPPA5/ESG1,SSEA-1, EpCAM/TROP1, SSEA-3, ERR beta/NR3B2, SSEA-4, ESGP, STAT3, F-boxprotein 15/FBXO15, Stella/Dppa3, FGF-4, SUZ12, FGF-5, TBX2, FoxD3, TBX3,GBX2, TBX5, GCNF/NR6A1, TERT, GDF-3, TEX19, Gi24/VISTA/B7-H5, TEX19.1,Integrin alpha 6/CD49f, THAP11, Integrin alpha 6 beta 1, TRA-1-60(R),Integrin alpha 6 beta 4, TRA-1-81, Integrin beta 1/CD29, TROP-2, KLF4,UTF1, KLF5, ZIC3, or L1TD1. Exemplary iPS markers include SSEA-4,Alkaline Phosphatase/ALPL, Oct-3/4 and Nanog.

In various embodiments of the method of recellularizing a tissue ororgan scaffold according to the disclosure, one or more antibodyproducts binding an endothelial cell marker, such as the exemplaryendothelial cell markers identified above, is/are used to facilitate orenhance the association or attachment of cells to the scaffold.Embodiments are also contemplated in which one or more antibody productsbinding an induced pluripotent stem cell marker, such as the exemplaryiPS markers identified above, is/are used to facilitate or enhance theassociation or attachment of cells to the scaffold. In yet additionalembodiments, a mixture of antibody products is contemplated for use inthe methods of recellularizing a tissue or organ scaffold, i.e., atleast one antibody product recognizing an endothelial cell marker incombination with at least one antibody product recognizing an inducedpluripotent stem cell marker.

The disclosure will be more fully understood by reference to thefollowing examples which detail exemplary embodiments of the disclosure.The examples should not, however, be construed as limiting the scope ofthe disclosure.

EXAMPLE 1 Material & Methods

The materials and methods disclosed in this Example are applicable tothe studies described in Examples 1-8.

Kidney Procurement

Ten human kidneys procured for transplant purposes but eventuallydiscarded for the reasons mentioned above, were processed. All organswere procured within the designated service area of our local organprocurement organization (Carolina Donor Services) and were refused byall local, regional, and national transplant centers. After exhaustionof the national list, kidneys from donors with research consent wereoffered for research purposes to the transplant team of the Wake ForestSchool of Medicine and processed at the Wake Forest Institute forRegenerative Medicine.

Kidney Preparation

Kidneys were received in sterile cold storage and placed in a sterilebasin containing ice and preservation solution used for shipment. Theaortic patch and the renal vein were prepared as for transplantpurposes. The renal artery was dissected circumferentially towards thehilum of the kidney. The renal vein was dissected circumferentially andsectioned at 2 cm from its origin. In the case of multiple arteries,these were reconstructed in order to create a single arterial inlet. Theexcess perinephric fat was trimmed from the kidney, leaving a triangleof fat at the lower pole. The perihilar fat and lymphatic tissue wasligated with 2-0 silk ties. Sixteen gauge intravenous catheters wereinserted into the ureter and the renal artery and secured with 2/0 silksuture to allow perfusion. Thereafter, the renal artery and ureter weretested for possible leaks and repaired with 6-0 Prolene figure-of-eightsutures. Because all kidneys had been biopsied at the upper pole at thetime of procurement, a renorrhaphy of the “wedge” defect was performedwith 4-0 PDS suture in a running fashion. Kidneys were finally placed onice until decellularization.

Decellularization—Kidney (One Protocol)

At room temperature, the angiocatheters previously inserted in the renalartery and in the ureter were connected to a pump (Masterflex L/Speristaltic pump with Masterflex L/S easy load pump head and L/S 16Gtubing, Cole-Palmer Instrument Co, Vernon Hills, Ill., USA) to allowcontinuous rinsing with different solutions, starting with distilledwater at a rate of approximately 12 ml/min for 12 hours (8,640 mLtotal), as previously described¹². Afterward, 0.5% sodium dodecylsulphate (SDS)-based solution was delivered at the same flow rate for 48hours (34,560 mL total) in both the renal artery and ureter. Finally,the kidneys were rinsed with phosphate buffer saline (PBS) for 5 days ata flow rate of 6 mL/min (43,320 mL total).

Decellularization—Kidney (Another Protocol)

Kidneys are received in sterile cold storage and placed in a sterilebasin containing ice and preservation solution used for shipment. Theaortic patch and the renal vein are prepared for research in the mannerprepared for transplant purposes. The renal artery is dissectedcircumferentially towards the hilum of the kidney. The renal vein isdissected circumferentially and sectioned at 2 cm from its origin. Theexcess perinephric fat is trimmed from the kidney, leaving a triangle offat at the lower pole. The perihilar fat and lymphatic tissue is ligatedwith 2-0 silk ties. Sixteen gauge intravenous catheters are insertedinto the ureter and the renal artery and secured with 2/0 silk suture toallow perfusion. Thereafter, the renal artery and ureter are tested forpossible leaks and repaired with 6-0 Prolene figure-of-eight sutures.Because all kidneys are biopsied at the upper pole at the time ofprocurement, a renorrhaphy of the “wedge” defect is performed with 4-0PDS suture in a running fashion. Kidneys are finally placed on ice untildecellularization.

At room temperature, the angiocatheters previously inserted in the renalartery and in the ureter were connected to a pump (Masterflex L/Speristaltic pump with Masterflex L/S easy load pump head and L/S 16Gtubing, Cole-Palmer Instrument Co, Vernon Hills, Ill., USA) to allowcontinuous rinsing with different solutions, starting with distilledwater at a rate of approximately 12 ml/min for 12 hours (8,640 mLtotal). Afterward, 0.5% sodium dodecyl sulfate (SDS)-based solution wasdelivered at the same flow rate for 48 hours (34,560 mL total) in boththe renal artery and ureter. After decellularization with SDS, 1000 mLof DNase solution (0.0025 w/w % DNase [Sigma-Aldrich, DN25] and 10 mmmagnesium chloride [Sigma-Aldrich, M4880] in 1× PBS at neutral pH) arere-circulated through the hrECMs overnight to allow digestion ofresidual DNA. Thereafter, hrECMs are rinsed with phosphate buffer saline(PBS) for 5 days at a flow rate of 6 mL/min (43,320 mL total). Finally,the resulting hrECMs are stored in PBS and, eventually, gamma-irradiatedto achieve sterilization.

Decellularization—Pancreas

First, the pancreata are procured with the duodenum and the spleenattached. Pancreata are received in sterile cold storage and placed in asterile basin containing ice and preservation solution used forshipment. For research purposes, the splenic artery and the superiormesenteric artery are prepared as for transplant purposes. Then, theduodenum is removed to allow identification of the pancreatic duct,which is “buried” in the duodenum. Likewise, the spleen is removed,after ligation and section of the distal segment of the splenic vein andartery, as well as of the short gastric arteries. The excessperipancreatic fat and lymphatic tissue are ligated with 2-0 and 3-0silk ties, and trimmed from the organ thereafter. Sixteen gaugeintravenous catheters are inserted into the splenic artery, the superiormesenteric artery and the pancreatic duct and secured with 3/0 silksutures to allow perfusion. Thereafter, the two arteries and the ductare tested for possible leaks and repaired with 6-0 Prolenefigure-of-eight sutures. The organ is finally placed on ice untildecellularization.

Decellularization takes place in the following steps:

1. Wash with 10% betadine for 10 minutes. In one embodiment, the organis put in a basin containing betadine in order to kill bacteria.

2. Wash with 10% Penicillin/Streptomycin for 10 minutes. Wash with 10%antibiotics for 10 minutes.

3. Rinse with 500 mL sterile PBS.

4. Place pancreas in container, connect lines and secure with silksutures. For example, one line goes into the pancreatic duct and thesecond one is split into two (the first one is connected to the superiormesenteric artery and the second one goes into to the splenic artery).

5. Pre-fill the container and prime the pancreas with PBS and Heparin(1% of 1000 U, 10 U/mL).

6. Seal the container and place in a refrigerator (4° C.).

7. Flush the pancreas with PBS and Heparin (1% off 1000 U, 10 U/mL) for15 minutes at a rate of 0.75 L/hr in each inlet (total PBS volume is187.5 ml for the pancreatic duct and 93.75 ml for mesenteric artery andsplenic artery).

8. Flush the pancreas with 1% Triton and 0.1% ammonium hydroxide in PBSfor 48 hours at a rate of 0.75 L/hr in each inlet (total volume is 36 Lfor the pancreatic duct and 18 L for the mesenteric and the splenicartery).

9. Flush the tissue or organ with PBS for 4 days at a rate of 0.375L/hour in each inlet.

10. Flush for 24 hours with PBS and 1% Penicillin/Streptomycin andGentamicin at a rate of 0.375 L/hour in each inlet.

11. Flush for 4 hours with DNase and 0.0025% magnesium chloride at arate of 0.375 L/hour in each inlet.

12. Flush for 12 hours with PBS at a rate of 0.375 L/hour in each inlet.

Bioscaffold Characterization Basic Histology

To assess cell and nuclear clearance as well as preservation of collagenfibrils, hematoxylin and eosin (H&E), Masson's trichrome (NewcomerSupply, Middleton, Wis., USA) and nuclear-specific4,6-diamidino-2-phenylindole (DAPI) staining were performed afterfixation in formalin 10%, paraffin embedding and sectioning. We expectednuclei and cells to be completely removed and collagen fibrils to bepreserved.

Immunohistochemistry (IHC)

Tissue slides were fixed in 4% paraformaldeyhyde, quenched forendogenous peroxidase in 70% Methanol plus 0.3% Hydrogen Peroxide,blocked with protein blocker (Dako, X090930) and incubated with primaryantibodies diluted in antibody diluent (Dako S0809) for 1 hour at roomtemperature. Tissue slides were washed in 3× PBS, incubated withsecondary antibodies for 30 minutes at room temperature, washed with 3×PBS and incubated with ABC reagent (Vector Elite ABC Reagent, VectorPK-7100). Target proteins detection was visualized by adding DAB reagent(Vector ImmPACT DAB Peroxidase Substratek, Vector SK-4105). Cell nucleiwere counterstained by hematoxylin.

Antibodies for Immunostaining

1) Primary antibodies for immunostaining included: Anti-HLA DR [LN-3](Abcam Ab49388, 1:50), Anti-HLA class 1 ABC[EMR8-5] (Abcam ab70328,1:50), Type IV collagen (Developmental Studies Hybridoma Bank, M3F7,1:100, which binds to the triple helical domain of collagen alpha1 andalpha2 chains), Anti Collagen IV [COL-94] (Abcam Ab6311, 1:50), Anticollagen IV alpha3 (Sigma SAB4500376, 1:50), COL4A4[G-12] (Santa Cruzsc-167524, 1:50), Anti-Agrin (Abcam Ab85174, 1:100), COL4A5 [H-234](Santa Cruz sc-11360, 1:50), COL4A6 [H-299] (Santa Cruz sc-134614,1:50), Nidogen-1/Entactin (R &D system AF2570, 1:50), Nidogen-2 (R &Dsystem AF3385, 1:50), Laminin alpha1 (LAMA1)(Lifespan BiosciencesLS-C25112/34991, 1:50), Laminin alpha5 (LAMAS) (Lifespan BiosciencesLS-C119651/28195, 1:100), Laminin a5-chain (Millipore MAB 1924, 1:100),Laminin (Millipore MAB1921, 1:50), Anti-integrin alpha3/beta1 [P1B5](Abcam Ab24696, 1:100).

2) Secondary antibodies included biotinylated goat anti-rabbit IgG(1:200) (Vector), biotinylated goat anti-mouse IgG (Vector),biotinylated goat anti-rabbit IgG (1:200) (Vector), biotinylated goatanti-mouse IgG (1:200) (Vector). Target protein detection reagentsincluded VECTASTAIN Elite ABC Reagent, R.T.U. (Vector PK-7100) andImmPACT DAB Peroxidase Substrate (Vector SK-4105), which were used forIHC staining. Positive staining (typically black) was the expectedresult.

Scanning Electron Microscopy (SEM)

A segment of decellularized kidney incorporating both medulla and cortexwas fixed in 2% glutaraldehyde in 0.1 M phosphate buffer and left for 24hours at 3° C. Following washing with 0.1 M phosphate buffer, 0.5 cm²pieces of tissue were sampled from both the medulla and cortex andcryoprotected in 25% sucrose, 10% glycerol in 0.05 M PBS (pH 7.4) for 2hours, then fast frozen in nitrogen slush and fractured at approximately−160° C. The samples were then placed back into the cryoprotectant atroom temperature and allowed to thaw. After washing in 0.1 M phosphatebuffer (pH 7.4), the material was fixed in 1% OsO⁴/0.1 M phosphatebuffer (pH 7.3) at 3° C. for 1.5 hours and washed again in 0.1 Mphosphate buffer (pH 7.4). After rinsing with distilled water, specimenswere dehydrated in a graded ethanol-water series to 100% ethanol,critical point dried using CO² and finally mounted on aluminum stubsusing sticky carbon taps. The material was mounted to present thefractured surfaces to the beam. The complete samples were opened andmounted to show the fractured surface, then coated with a thin layer ofAu/Pd (approximately 2 nm thick) using a Gatan ion beam coater. Imageswere recorded with a Jeol 7401 FEG scanning electron microscope.

DNA Quantitation

The DNA content of fresh and decellularized kidney was quantified usingthe tissue DNA isolation kit (PureLink Genomic DNA MiniKit, Invitrogen)according to the manufacturer's instructions. Briefly, the samples weredigested overnight using Proteinase K and a digestion buffer. Uponremoval of RNA, DNA samples were isolated by spin column-based nucleicacid purification and the extracts were characterizedspectrophotometrically (NanoDrop 1000; Thermo Scientific). Opticaldensities at 260 nm and 280 nm were used to estimate the purity andyield of nucleic acids, which were quantified on the basis of 280 nmabsorbance.

Glycosaminoglycan and Collagen Quantification

The sulfated glycosaminoglycan (GAG) content of fresh and decellularizedkidneys was quantified using the Blyscan GAG Assay Kit (Biocolor, UK).In brief, 30 mg of minced wet tissue were digested in papain-containingdigestion buffer for 18 hours at 65° C. Aliquots of each sample weremixed with 1,9-dimethyl-methylene blue dye and reagents from the GAGassay kit. The absorbance at 595 nm was measured using a microplatereader (Tecan Infinity) and compared to standards made from bovinetracheal chondroitin-4-sulfate to determine the absolute GAG content.

The collagen content was measured indirectly through measurement ofhydroxyproline residues using the QuickZyme total Collagen Assay(QuickZyme, Biosciences). Five mg of tissue was acid-hydrolyzedovernight with 6 M HCl at 95° C., following assay as described by themanufacturer. Hydroxyproline content resulted in a chromogen with anabsorbance maximum at 570 nm, which was measured using a microplatereader (Tecan Infinity). The absolute collagen content was determined bycomparison to standards from hydrolyzed collagen (1200 μg/ml in 0.02Macetic acid).

Chicken Chorioallantoic Membrane (CAM) Angiogenic Assay

The CAM was performed essentially as previously described [28, 29].Fertilized chicken eggs (Henry Stewart and Co., UK) were incubated at37° C. and constant humidity. At 3 days of incubation an artificial airsac was created by aspirating 2 ml of albumin from the acute end of theegg using a 21-G gauge needle. An oval window of approximately 3 cm indiameter was opened in the shell with small dissecting scissors toreveal the embryo and CAM vessels. The window was sealed with tape andthe eggs were returned to the incubator for a further 5 days. At day 8of incubation, 1 mm diameter samples of kidney acellular ECM (3 samples)or polystyrene (three samples) as a negative control, were placed on theCAM between branches of the blood vessels. Grafts were examined dailyuntil 7 days after placement wherein they were photographed with astereomicroscope equipped with a Camera System (Leica) both in ovo andafter removal and pinning in a silicone rubber-coated dish, to quantifythe blood vessels surrounding the matrices. The number of blood vesselsless than 10 μm in diameter converging towards the placed tissues wascounted blindly by assessors (scaffold n=3, polystyrene n=3, assessorsn=3).

Pressure Measurement

To test the compliance of vascular network of the human renal ECMscaffolds, native and decellularized kidney samples were perfused withwater through the cannulated renal artery using a peristaltic pump(Masterflex 7523-70, Cole-Parmer) and pulse dampener (Cole-Parmer) tocreate steady flow. Pressure was measured and recorded using a MillarMPC-500 Mikro-Tip pressure transducer with MPVS-400 signal conditioninghardware (Millar Instruments Inc.) and integrated data acquisitiontechnology (Powerlab, ADInstruments). The pressure probe was insertedinto the renal artery and pressure was recorded at flow rates rangingfrom 10 ml/min to 30 ml/min in 5 native and 6 decellularized kidneys.All probes were calibrated to mmHg before each use.

EXAMPLE 2 Human Kidney Decellularization

Perfusion of SDS-contained buffer successfully decellularized humankidneys discarded from transplantation. DNA confirmed this observationand showed that approximately 95% of DNA was removed in comparison withthe native organ (12.27±0.7 vs. 274.7±4.7, p<0.0003). On the other hand,measurements of glycosaminoglycans (GAGs) demonstrated a decrease from0.89±0.18 μg/mg to 0.55±0.21 μg/mg, equal to a reduction to 60% of thefresh value (p=ns). Similar results were observed for the amounts ofcollagen remaining in the acellular ECM scaffolds.

EXAMPLE 3 Histological Characterization of the Human Renal ECM Scaffold

H&E staining of the decellularized kidney tissue showed pinkeosinophilic staining typical of collagen while no basophilic stainingindicative of cellular nuclear material was detected (FIGS. 2A, B). Thescaffolding architecture of glomeruli, vessels and tubules waspreserved. Notably, our decellularization protocol allowed cellclearance despite severe glomerulosclerosis and interstitial fibrosis.FIG. 2B shows an acellular glomerulus at an advanced degree of ischemiccollapse with a remnant of the corrugated glomerular basement membraneof the collapsed tuft and surrounded by the thickened and multilayeredBowman capsule basement membrane. Masson's trichrome stain confirmedthese results, showing a homogenous blue staining of the ECM consistentwith collagen (FIGS. 2C, D). This indicates that the disclosed methodallows detergents to reach all cellular compartments despite advancedfibrosis. Methenamine silver staining was performed in order to assesspreservation of glomerular basement membrane (GBM), which is anessential structure of renal ECM scaffolds situated between endothelialcells and podocytes. This membrane derives from the fusion of theendothelial cell and podocyte basal laminas and represents the basallaminal portion of the glomerulus that performs the actual filtrationthrough the filtration slits between the podocytes, separating the bloodon the inside from the filtrate on the outside. The methenamine silverstaining revealed the presence of a well preserved GBM (FIGS. 2E-H),which was often thick because of severe glomerulosclerosis in thesediscarded kidneys. To further assess the decellularization efficiency,cell nuclei staining was performed using DAPI, and confirming successfulclearance of cells and nuclear material (FIGS. 2I-J).

EXAMPLE 4 Removal of Antigenic Markers

The HLA system is the major histocompatibility complex in humans. ABCantigens represent the human major histocompatibility complex class I,while DR antigens are members of class II. They are heavy chainreceptors expressed in nearly all cells whose role is to help the immunesystem distinguish the body's own proteins from non-self proteins. HLAclass I antigens (A, B, and C) present peptides from inside the cell,while HLA class II (DR) present antigens from outside of the cell toT-lymphocytes [32]. Immunostaining for HLA-ABC and HLA-DR showed thatthe antigens were completely removed upon decellularization, whereasthey strongly present in the native kidneys (FIGS. 3A-D). This findinghas a significant immunological implication, as it indicates that humanrenal ECM scaffolds may have only minimal if not negligibleimmunogenicity, possibly limited to a non-specific inflammatoryresponse. Integrin α3/β1 are transmembrane heteromeric receptors thatmediate interactions between cells and ECM, are highly expressed in theglomerulus of the kidney, and are key players in glomerularmorphogenesis and maintenance of glomerular filtration barrierintegrity. Similar to the efficient removal of the HLS-ABC and HLA-DRantigens, the decellularization protocol efficiently removed integrinα3/β1 in the acellular human renal ECM scaffolds (FIGS. 3E, F).

EXAMPLE 5 Immunohistochemical Characterization of the Human ECM RenalScaffolds

Type IV collagen (Coll IV) is the major structural component of GBMtogether with laminins, proteoglycans and nidogen. Laminins areubiquitous basement membranes. The most important receptors for lamininα5 are the integrin family. Agrin is the major heparan sulfateproteoglycan of the GBM playing a key role in the renal filtration andcell-matrix interactions. Nidogen 1 and 2 are both dumbbell-shaped,virtually ubiquitous basement membrane proteins, binding to the shortarm of laminin γ1 as well as to Coll IV. Staining for Coll IV, with 2different anti-Coll IV antibodies, showed preservation of Coll IV in theGBM (FIGS. 4A-D). Laminin α5 was also preserved after decellularizationof the human kidneys, and its preservation is important for successfulcell attachment on the renal scaffolds (FIGS. 4E-F). Agrin and nidgenswere also well preserved (FIGS. 4G-J).

EXAMPLE 6 Ultrastructural Analysis

SEM confirmed the successful clearance of the cellular compartment ofhuman kidneys (FIGS. 5C-F). No cells or nuclei could be observed, whilethe remaining acellular ECM scaffold maintained the ultrastructuralcharacteristics of the native tissue (FIGS. 5A-B). The three-dimensionalarchitecture of the kidneys was well preserved including all of theessential structures—namely, glomeruli, tubules and vessels of anyhierarchical level, which were well represented and evident. Lowermagnification images of a single renal corpuscle show the integrity ofthe glomerulus with its winding blood capillary loops (FIGS. 5C-D). Thecapillary wall is intact and no podocytes are present on the outersurface (FIG. 5C). SEM view of the microdissected glomeruli showed theconvoluted GBM and the vascular pole where the afferent arterioleintegrates with the glomerulus (arrow) (FIG. 5D). The surface of theGBM, under higher magnification, is regular and no cellular elementscould be observed (FIG. 5E). The internal surface of the parietal layerof the Bowman's capsule appeared like a flat, regular surface where noepithelial cell could be detected. A transversal section of the sampleof renal ECM gave a honeycomb-like image of the complex network ofarterioles and tubules, which composes the nephron structure in themammalian kidney and was well preserved in the decellularized tissue(FIG. 5F).

EXAMPLE 7 Induction of Angiogenesis

The chorioallantoic membrane (CAM) of chicken embryos is a highlyvascularized structure normally adherent to the egg shell andresponsible for gas exchange, and thus, used in in vivo assay todetermine the angiogenic/antiangiogenic activities of biomaterials andother compounds. Macroscopic observation of the CAM at day 0 showed therenal ECM scaffold on top of the allantoic vessels beneath, but withoutvessels penetrating the scaffold (FIG. 6B). Five days later the scaffoldhad changed color to a yellowish hue, was well-adherent to the CAM andnew vessels of the CAM had begun penetrating renal ECM scaffolds (FIG.6C). At 7 days, the ECM scaffolds were completely enveloped by the CAMand the vessels, which formed an organized network around the scaffolds(FIG. 6D). The pro-angiogenic effect of the renal ECM scaffolds wasquantified at day 0, 5 and 7 in a blinded fashion. Human renal ECMscaffolds induced the onset of a significantly higher number of vessels,both from longitudinally (i.e., from time 0 to day 7) and versus thecontrol. At day 0 (time of transplantation) no vessels growing towardsthe implanted tissues were observed. At day 5 the renal ECM scaffoldssignificantly stimulated the growth of converging allantoic vesselscompared to day 0 (p=0.007). A further increase was observed at day 7,with the number of converging allantoic vessels significantly higherwhen compared to day 0 (p<0.0001) and to the control polystyrenemembranes at the same time-point (p<0.0001; FIG. 6A).

EXAMPLE 8 Resistance to Vascular Pressures

The average renal artery pressure in native kidney ranged from about32-44 mmHg at 10 ml/min and increased linearly to about 50-94 at mmHg at30 ml/min (R²=0.99) (FIG. 7). The pressures measured for the acellularrenal ECM scaffolds ranged from about 10-195 mmHg at 10 ml/min andincreased linearly to about 17-50 mmHg at 30 ml/min (R²=0.91) (FIG. 7).One-way ANOVA determined that a statistically significant differenceexists between native and acellular kidney artery pressure (p<0.05).Post-hoc analysis using Fisher's least significance difference (LSD)correction was conducted and determined that statistically significantdifferences (p<0.05) were observed between native and acellular kidneypressure at 20 ml/min, 25 ml/min, and 30 ml/min (FIG. 7). Although therewas a large degree of variability in samples, possibly due the varyingdegree of vascular damage (arteriolo- and arterio-sclerosis, vascularhyalinosis) present in the renal grafts, the linear increase inpressures indicates good preservation of the vasculature in the renalECM scaffolds, with patent and resilient vasculature.

REFERENCES—FOR EXAMPLES 1-8 ONLY

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The data presented in Examples 1-8 show that discarded human kidneys canbe successfully decellularized with the resulting renal ECM scaffoldsmaintaining (a) their three-dimensional architecture at the macro andmicro levels, (b) vascular patency and (c) biological function. From aclinical transplant perspective, the possibility of utilizing discardedhuman kidneys as source of organs gives hope to the myriad of patientswho are currently waiting for a new kidney and have to face longerwaiting times because of the increasing disparity between supply anddemand. The catastrophic consequence represented by the increase inwaiting list mortality is fueling efforts to meet the urgent need toidentify new organ sources. Organ bioengineering and regeneration holdsthe promise to meet this need and justifies investments and efforts inthis direction.

EXAMPLE 9 Whole Organ Engineering—Antibody Conjugation toRe-Endothelialize Kidney Scaffolds

Consistent with observations made above, end-stage renal disease (ESRD)is a major problem in the United States and around the world. Accordingto a report by the National Kidney & Urologic Diseases InformationClearinghouse (2009), more than 870,000people in the United States wereundergoing treatment for ESRD. Current treatments are limited tolife-long dialysis and/or renal transplantation. While dialysis canreplace the filtration function of the kidneys by removing certaintoxins from the patient's blood, it cannot replace many of the otherfunctions of the kidneys, such as production of erythropoietin andvitamin D; resulting in complications such as anemia, malnutrition, andeventually requires kidney transplantation1. Therefore, kidneytransplantation is currently the only definitive solution for thetreatment of ESRD. However, due to a critical shortage of availabledonor kidneys, physicians and scientists have sought for novel solutionsto this problem.

Whole organ engineering techniques based on decellularization of organsand recellularization of the resulting collagen-based matrix wouldprovide an effectively inexhaustible supply of organs to replace thosesuffering from disease or injury. The first step of this technique isthe “decellularization” of native organs. During the decellularizationprocess, primary immunogenic factors such as cellular proteins and DNAcomponents are removed from the organ so that foreign body responses canbe reduced after transplantation. Decellularization methods aretypically based on perfusion of the organ through existing vascularsystems with detergents and other agents. These perfusion-baseddecellularization protocols allow efficient removal of all native cellsand residual DNA, while maintaining the structural integrity of theorgan's extracellular matrix. Subsequently, the resulting acellularmatrix needs to be repopulated with functional organ-specific cellpopulations (“recellularization”). Available cell sources fortarget-specific organs can be seeded onto the decellularized matrix andallowed to organize into functional components of the organ. Ideally,engineered whole organs should contain an intact three-dimensionalcellular architecture as well as functional vasculature that is composedof appropriate cell types to achieve organ function aftertransplantation.

The field of whole-organ tissue engineering is still in its infancy. Onemajor challenge for long-term in vivo success of bioengineered organs isvascular patency. In the absence of complete endothelial reseeding ofvascular matrices, significant thrombosis is likely to occur within thevasculature of the scaffold, thus rendering the recellularized constructnonfunctional. To address this issue, we have developed an endothelialcell-seeding method that permits effective endothelial cell coating ofthe vascular walls of decellularized porcine kidney scaffolds. Acellularrenal matrices were processed from normal pig kidneys using adecellularization technique incorporated herein by reference¹³. Severalendothelial cell-seeding methods were tested that have been described inother studies^(2,4,5,8). Through optimization of these cell-seedingprotocols, a combined cell-seeding method was developed for acellularkidney scaffolds that is composed of static and ramping perfusionseeding. The efficiency of re-endothelialization on the blood vessels inthe kidney scaffold was determined by histological characterization andimaging studies of the re-endothelialized vascular tree, as well as invitro functionality being tested by blood perfusion. In addition, asurface modification method was developed to reinforce endothelial cellattachment onto renal vasculatures via CD31 antibody conjugation. Theeffectiveness of antibody conjugation to mediate re-endothelializationin vivo was evaluated through implantation (FIG. 23).

Decellularization of porcine kidney and endothelial cell seeding ofacellular kidney scaffolds

Decellularization of porcine kidney was processed using ahigh-throughput system as described¹³, and that description isincorporated herein by reference. In brief, whole kidneys were perfusedwith heparin and then 0.5% sodium dodecyl sulfate (SDS)inphosphate-buffered saline (PBS) for 36 hours, followed by a rinse withPBS, enzymatic treatment with 0.0025% DNase (Sigma-Aldrich, St. Louis,Mo.) for 12 hours, followed by another rinse with PBS. Subsequently,decellularized kidneys were sterilized via gamma irradiation at 1 MRad(10,000 Gy) prior to cell seeding. Vascular endothelial cells expressingGFP protein (MS1) were used in this study⁵. Four experimental groupswere tested in order to find an optimal cell seeding condition. Theseinclude (1) perfusion cell seeding (100×10⁶) at a constant rate of 20ml/min (n=2), (2) static cell seeding (50×10⁶) followed by constantperfusion seeding (50×10⁶) at 20 ml/min (n=1), (3) ramping perfusioncell seeding (100×10⁶) (n=2), and (4) static cell seeding (50×10⁶)followed by ramping perfusion (50×10⁶) (n=3). For static cell seeding, 5ml of 10×10⁶/ml MS1 cells suspended in DMEM-high glucose medium(Invitrogen Life Technologies, Carlsbad, Calif.) supplemented with 10%FBS (Invitrogen Life Technologies) was infused into the renal artery(2.5 ml) and vein (2.5 ml) simultaneously. The seeded cells were thenallowed to attach for 2 hours, followed by another 2 hours after a180-degree angle rotation of the kidney scaffold from the originalposition to allow for homogenous attachment of the cells in an incubatorat 37° C., supplemented with 5% CO₂. Following the static seeding, thekidney scaffold was connected to a bioreactor system and then perfusedat 2 ml/min with 10% FBS DMEM medium until ramping perfusion seedingbegan. The ramping perfusion was initiated at a rate of 2 ml/min andgradually increased to 5, 10, and 20 ml/min at 10-12 hour intervals, andmaintained at 20 ml/min. All kidney scaffolds received a total ofapproximately 100×10⁶ cells and were cultured for 3 days in thebioreactors prior to analysis. For the constant perfusionseeding,100×10⁶ MS1 cells were suspended in the bioreactor medium, andthe scaffold was continuously perfused at a constant flow rate of 20ml/min. Almost all of the seeded cells were retained within thescaffold. This was confirmed by examining the returned perfusate in thebioreactor container. Only a few cells were visible under a 10×microscope. After bioreactor perfusion culture, the MS1-seeded scaffoldwas fixed in 10% formalin, embedded in OCT compound, and processed forhistological analysis (H&E and DAPI nuclear staining).

Characterization of Re-Endothelialization on Kidney Scaffold

Seeding efficiency was determined by histomorphometric analysis usingH&E stained images of the re-endothelialized vessels of different sizes(large for renal artery and vein, intermediate, small) within thescaffold. The number of blood vessels that had been re-endothelialized(greater than 50%coverage in each vessel) and/or filled with MS1 cells(greater than 50% occupancy of vessel spaces) were counted and scoredfor analysis (Table 1).

TABLE 1 Static + Static + Constant constant Ramping ramping Celladhesion perfusion perfusion perfusion perfusion Artery and vein + + ++++ Intermediate-sized + + ++ +++ blood vessels Small-sized blood + + +++++ vessels in cortex Cell clogging ++ +++ + + Criteria for scoring:0-30% (+), 31-60% (++), 61-100% (+++).

Vessel size greater than 100 μm was considered as intermediate, and lessthan 100 μm as small. Re-endothelialized renal artery and vein wereobserved under scanning electron microscopy (SEM; ModelS-2260N, HitachiCo. Ltd., Japan). Briefly, the sample was sputter-coated with gold(Hummer™ 6.2, Anatech Ltd, Denver, N.C., USA) to a thickness of 10-15nm. Images were acquired using an environmental SEM operating at anaccelerating voltage of 20 kV with a 10 cm working distance. Forre-endothelialized vascular tree imaging, formalin-fixed blood vesselbranches were isolated and visualized with a fluorescentstereomicroscope (Leica, Germany) equipped with a GFP protein detectionfilter.

In vitro Functional Testing

To determine the functionality of re-endothelialized kidney vasculature,seeded scaffolds were perfused with freshly collected heparinized pigblood for 30 minutes up to 100 ml/min of flow rate. To more closelyrepresent an in vivo environment following implantation, the perfusionrate was increased up to 100 ml/min, which is the maximum rate the pumpcan generate. The blood perfusate was prepared by diluting whole bloodwith 3:1 ratio in Kreb's bicarbonate buffer. Unseeded scaffolds wereused as controls. To determine platelet adhesion on the scaffolds,frozen sections were stained with an anti-integrin αIIb antibody(SantaCruz Biotechnology Inc., Santa Cruz, Calif.) and Alexa647-conjugated rabbit anti-goat antibody (Invitrogen Life Technologies).Images were visualized with an inverted microscope (Nikon Eclipse TE2000-U) with an argon laser confocal microscope system (Laser Drive Inc,Gibsonia, Pa.) facilitated with MAG Biosystems software. The numbers ofstained platelets were counted in 3 randomly selected images fromdifferent sized vessels. The counts were averaged, and then representedas mean±SD.

Cell Detachment Study Under Flow Conditions

A polydimethylsiloxane (PDMS)-based microfluidic device was constructedto conduct cell detachment study under shear flow. The molded portion offlow chamber was made from Sylgard 184 silicone elastomer kit (DowCorning, Midland, Mich.) according to manufacturer's instructions. Afterpreparation of the chamber, 20 G needles were inserted into the inletand outlet and tubing was used for connecting the inlet to syringes. Theinterior surface of the chamber was coated with type I collagen (ElastinProducts Co. Inc., Owensville, Mo.) by filling the chamber with a 0.05%collagen solution dissolved in 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP,Sigma). Rat anti-mouse CD31 antibody (BD sciences, Franklin Lakes, N.J.)was conjugated to the collagen-coated chamber by filling the chamberwith a solution of 1-Ethyl-3-[3-dimethylaminopropyl]carbodiimidehydrochloride (EDC) (Sigma-Aldrich) and N-hydroxysuccinimide esters(NHS) (Sigma-Aldrich) in PBS buffer for 30 minutes, followed by reactingwith a 100 μg/mL antibody solution for 2 hours. Control chambers werecoated with collagen, crosslinked with DC/NHS, and then treated with PBSfor 2 hours. Collagen coating and CD31 antibody conjugation on thesurfaces of chamber slides was confirmed with coomassie blue stainingand treatment with Alexa588 goat anti-rat antibody, respectively. Theconjugated antibody was semi-quantified by imaging with a fluorescentmicroscope and measuring the fluorescent intensity using Image Jsoftware. To examine the efficacy of antibody conjugation on reducingcell detachment from the surfaces, the flow chamber experiment underdynamic flow conditions (2 to 100 ml/hour) was performed. For the celldetachment study, MS1 cells were allowed to pre-adhere on the surfacesby placing the cell suspension solution (10⁵ cells/mL) at staticconditions for 10 minutes before starting flow through the chamber. Aduration of 10 minutes was chosen for this experiment, based on previousstudies on cell adherence and detachment¹⁴. A syringe pump was used toflow 3% w/v BSA/media through the chamber at a rate of 2 mL/hour for 10minutes to remove non-adherent cells. The flow rate of media through thechamber was increased sequentially to 10, 20, 50, and 100 mL/hour witheach flow rate run for 3 minutes. Continuous video of the cells wasobtained during the flow. The number of cells on the surface was countedafter exposure to each flow condition. All values were normalized to thenumber of cells after 2 mL/hour flow. Cell numbers were counted usingImage J software, averaged, and represented as mean±SEM.

Bioreactor Culture

Perfusion bioreactors were designed and constructed specifically forporcine kidneys, and consisted of a glass chamber for the kidney, withmedia circulation via platinum-cured silicone tubing (Cole-Parmer, L/S15) and a pulse dampener to maintain constant flow using Cole-ParmerDigital Drive peristaltic pumps. Bioreactors contained 2 L of growthmedium (DMEM-high glucose with 10% fetal bovine serum, 1%antibiotic/antimycotic supplement) for each cell-seeding method and weremaintained in humidified conditions at 37° C., supplemented with 5% CO₂.Media was perfused directly into the kidney via the renal artery and,during the culture, media was constantly stirred via a magnetic stirrer.

In vivo Implantation Study

Yorkshire pigs (female, 40-50 kg, 3-6 months) were used for the in vivoanimal study. All surgical procedures were performed in accordance withthe Institutional Animal Care and Use Committee (IACUC) approvedprotocols. The animals received two different groups of implantation:(1) 2 hours (re-endothelialized kidney without antibody conjugation,n=3animals) and (2) 4 hours implantation (re-endothelialized kidney withCD31 antibody conjugation, n=3 animals). To prepare MS1-seeded kidneywith antibody conjugation, CD31 antibody was conjugated to nativecollagen present in the decellularized kidneys¹³ using the same reactionprotocol used for the flow-chamber study. MS1 cell seeding on theantibody-conjugated scaffold was performed as described above(endothelial cell seeding method). The implantation protocol was similarto that developed in our previous study¹⁵. Briefly, animals were placedin a supine position, the surgical site was disinfected, and anintravenous catheter was placed to allow for proper fluid replacementand drug delivery during surgery, while monitoring cardiopulmonaryfunction. Under general anesthesia, a midline abdominal incision wasmade through the skin and muscle. The abdominal aorta and inferior venacava just above the iliac bifurcation were identified and dissected fromthe surrounding tissue. After placing a large Satinsky clamp on therecipient's inferior vena cava and aorta, the artery and vein of thekidney scaffold were anastomosed to the aorta and inferior vena cava inan end-to-side fashion using double-armed 6/0 prolene sutures. Afterreperfusion, the blood flow through the artery and vein of the implantedscaffold was monitored with Color Doppler ultrasonography (AcusonSequola 512, Simens).

Characterization of the Harvested Kidney

At 2 and 4 hours after implantation, patency of vascular tree in theharvested kidney scaffold from the euthanized animals was examined byradiographic fluoroscopy using a Siemens SIREMOBIL CompactL C-arm(Siemens, Munich, Bavaria, Germany). Conray (iothalamatemeglumine)(Mallinckrodt Inc., St Louis, Mo.) contrast agent was injected throughthe renal artery, followed by histological (H&E staining) andimmunohistochemical (platelet immunostaining) examination. To obtainpercentage of endothelialization by MS1 cells on the vasculatures, threedifferent images of medulla and cortical regions were taken (H&E andGFP). GFP+MS1-endothelialization was based on the criteria (greater than50%coverage in each vessel with less than 50% occupancy of vesselspaces). For quantification of platelet adhesion, αIIb+plateletfluorescent intensity was obtained from randomly selected images frommedulla (three sites)and cortex (nine sites). Fluorescence intensity wascalculated from the values [equal to (total fluorescenceintensity/area)] of each fluorescence image using Image J software. Thedata was represented as mean±SD.

Statistics

Two-way ANOVA statistical analysis and the Bonferonni post hoc test wereused for the flow chamber experiments. Student t-test analysis was usedfor other statistical analyses. Differences were considered significantat P<0.05.

Development of an Effective Cell Seeding Method: Combining Static andRamping Perfusion Cell Seeding

The first series of experiments were designed to identify and optimizean efficient technique for the successful cell seeding andre-endothelialization of the blood vessels using the MS1 endothelialcell line within an acellular kidney scaffold. A flow rate of 20 ml/minwas selected as the maximum perfusion rate, based on our previouscell-seeding experiments. Perfusion beyond this rate showed failure ofcell attachment in the scaffold vessels. For ramping perfusion cellseeding, cell suspension was perfused through the kidney vasculature ata rate of 2 ml/min, and gradually increased up to 20 ml/min. Gradualincrease in flow rate was employed to prevent cell clogging within thesmall capillary structures. Among the cell seeding methods tested,including constant perfusion, static+constant perfusion and rampingperfusion, the combination protocol of static and ramping perfusion wasshown to be the most effective method (FIG. 17 and Table 1) and, thus,this method of seeding was used for all other experiments in this study.We examined whether seeding of MS1 cells using the combined static andramping perfusion provides a homogenous distribution and functionalalignment of the MS1 cells within the vasculature of the kidneyscaffold. Following cell seeding, a vascular tree branching from therenal artery was isolated from the kidney parenchyma (FIG. 18 a) andvisualized with a fluorescent stereo-microscope in order to detectGFP-expressing MS1 cells (FIG. 18 b). The MS1-seeded vascular treestructure [EC(+)] showed an increase in GFP-derived fluorescentintensity compared to the unseeded matrix [EC(−)] (FIG. 18 b).Immunohistochemistry using CD31, vascular endothelial cadherin (VE-cad),and CD146 antibodies confirmed that the attached MS1 cells maintainedendothelial phenotypes on the renal artery when compared to that on thenormal culture condition (FIG. 24). Furthermore, fluorescence imaging ofparenchymal surfaces in the[EC(+)] kidney clearly showed capillary-likestructures on the cortical surface, indicating that viableGFP-expressing MS1 cells infiltrated the parenchymal cortex (FIG. 18 c).Scanning electron microscopy (SEM) confirmed endothelial cell alignmentin the vasculature (FIG. 18 d). Homogenous re-endothelialization ofdifferent-sized vasculatures was confirmed in different regions of renalparenchymal tissues (FIG. 25). These results showed that MS1 cells wereuniformly attached and well-organized on the vascular walls along thedirection of perfusion.

In vitro Functional Assessment of Re-Endothelialized Kidney Scaffolds

To determine whether the seeded kidney scaffolds are functional,heparinized pig blood was perfused through the artery. The functionalre-endothelialization of vasculatures was expected to decrease plateletadhesion on re-endothelialized renal scaffolds during blood perfusionand to maintain blood flow through the entire renal parenchyma. Bloodinfused through the artery of MS1-seeded kidney scaffolds showedsignificant outflow through the renal vein, indicating that endothelialcell coverage of the vasculature is effective and sufficient to supportblood flow throughout the entire kidney scaffold. In contrast, no bloodoutflow was observed through the renal vein of the EC(−) kidneys.Macroscopic images and H&E staining of the blood-perfused kidneyscaffolds confirmed the expectation that clotting was causing theblockage of blood flow in the EC(−) group. Gross images of perfusedEC(−) kidney scaffolds clearly showed multiple areas of thrombosis onthe parenchymal surfaces (FIG. 26). In contrast, EC(+) scaffolds showeda well-preserved endothelial lining without noticeable blood clots. Todemonstrate the effects of re-endothelialization, unseeded and seededkidney matrices perfused with blood, were immunostained with integrinαIIb antibodies. A minimum expression of integrin αIIb+platelets wasobserved on the re-endothelialized renal arteries, whereas high levelsof platelet accumulation were found in the EC(−) kidneys (FIG. 19 a).Quantitatively, the artery in EC(+) kidneys significantly reducedplatelet adhesion 4-5 fold compared to EC(−) kidneys (FIG. 19 b, P<0.05)Likewise, intermediate- and small-sized blood vessels lined with GFP+MS1endothelial cells showed no significant platelet adhesion, while strongplatelet-derived fluorescence was observed in the EC(−) kidneys (FIGS.19 c, e). The quantification of platelet adhesion by measurement offluorescence intensity showed a significant decrease in plateletadhesion in the seeded scaffolds (FIGS. 19 d, f). To examine whetherseeding of MS1 cells onto kidney scaffolds can effectively prevent bloodclots in vivo, MS1-seeded kidney scaffolds (FIG. 20 a) were implanted ina pig by arterial and venous anastomosis (FIG. 20 b). Followingimplantation, maintenance of blood flow was confirmed in both the renalartery and vein at 1 hour by Doppler Ultrasound (FIG. 20 c). Uponretrieval at 2 hours, the explanted kidney showed blood clots within thelarge blood vessels (FIG. 20 d) and occlusion of vascular network wasconfirmed by angiography. Several areas showed extravasation of contrastmedium in the parenchyma (arrows, FIGS. 20 e, f). Histological analysisof the explanted kidney scaffolds showed that most of the vasculaturethroughout the parenchyma in the implanted kidney failed to retain MS1cell coverage on the vascular walls, indicating that the failure ofvascular patency is due to a detachment of endothelial cells afterimplantation (arrows, FIGS. 20 g, h). Detachment of endothelial cellswas increased with implantation time increasing from2 to 4 hours. By 4hours, re-endothelialized kidney scaffolds had no measurable blood flowin the renal artery and vein (as confirmed by ultrasound imaging andangiography), which is thought to be due to an apparent increase inthrombi located inside the renal vasculature where detachment occurreddue to high vascular pressure and flow.

Effect of Antibody Conjugation on MS1Cell Detachment (in vivo) andEndothelial Integrity After Implantation

To address the apparent detachment of MS1 cells after implantation invivo, cell specific antibody was employed to strengthen the binding ofendothelial cells expressing CD31. Conjugating CD31 antibody (Ab) to thedecellularized vascular matrix was expected to improve MS1 cellattachment during cell seeding as well as cell retention underphysiological flow conditions. To test this expectation, CD31 Ab wasconjugated onto a collagen pre-coated surface of a microfluidic flowchamber system (FIG. 21 a). Conjugation of CD31 Ab was confirmed using afluorescent-labeled secondary antibody to the CD31 antibody (FIGS. 21 b,d). MS1 cells were then placed in the collagen-coated flow chambers±CD31Ab, and allowed to adhere to the surface for 10 minutes prior toassessing cell detachment in response to a stepwise increase in flow(2-100 ml/hour). There was no detectable endothelial cell detachment inthe Ab(+)-conjugated surfaces, whereas there was a significantendothelial cell detachment in the untreated surfaces at all flow ratestested (FIGS. 21 e, f, P<0.00001, n=4). These results indicate that CD31antibody conjugation onto the kidney vascular scaffold improves MS1 cellretention under physiological flow conditions. To determine whether CD31antibody conjugation could improve MS1 cell retention throughout therenal vascular scaffold in vivo, conjugated scaffolds were implantedinto pigs and exposed to physiological blood flow for 4 hours, afterwhich the implanted kidney construct was explanted and angiography wasperformed to assess vascular patency. Angiography results showed intactand patent vascular trees of parenchyma, which were withoutextravasation of host blood cells in the[Ab(+)] scaffolds during theentire period (FIG. 22 a), confirming the effectiveness of antibodyconjugation in the kidney scaffold. This functional outcome wassupported by the histomorphological analyses showing that CD31 antibodyconjugation significantly maintained endothelialization of intermediate-and small-sized vasculatures (FIG. 22 b). Moreover, the endothelium ofAb-conjugated constructs effectively prevented platelet adhesion (FIGS.22 c, d).

Whole-organ engineering using decellularized organ matrices provides thepotential to address the limitations of current transplantation. Onechallenge that must be addressed is to develop a method to preventthrombosis and maintain continuous blood flow into the engineeredorgan¹⁶. In this regard, re-endothelialization of scaffold vasculaturehas become a promising solution. Although approaches have been employedto establish endothelial cell coverage of vascular trees withindecellularized organs, complete and uniform coverage could not beachieved, thus resulting in the failure of long-term patency invivo^(2-5,8). Disclosed herein is an approach that has developed andoptimized a cell-seeding method that results in uniform coverage ofendothelial cells across different sizes of blood vessels within thekidney scaffold. More importantly, retention of seeded endothelial cellscan be maintained with the conjugation of CD31 antibody todecellularized vascular matrix prior to seeding. In this study severalcell seeding methods were tested; however, these techniques resulted inlimited or no cell attachment in the intermediate- and large-sized bloodvessels with cell “clogging” within the small capillaries in the renalcortex, indicating a need for improved cell seeding methodology. Assuch, the static and ramping-perfusion seeding methods, which resultedin uniform endothelial cell coverage within the vascular matrix of therenal scaffold, were combined.

Vascular patency as a function of endothelial cell-covered vessel wasdemonstrated by infusion of whole blood into the artery of kidneyscaffolds. To demonstrate the function of endothelial cellcovered-vessels, heparinized pig blood was infused into the renal arteryof the endothelialized scaffold. Blood perfused through the artery ofcell-seeded kidney scaffolds showed outflow through the renal vein.Moreover, the vasculatures within the scaffold maintained stableendothelial cell adherence under high flow conditions (100 ml/min) andshowed minimal expression of platelet adhesion, indicating thatendothelial cell-covered vasculature is effective and sufficient tosupport blood flow throughout the entire kidney scaffold. To test thefeasibility of implantation of engineered kidney scaffolds in vivo, asurgical technique previously developed in pigs was used, and theexperiment showed the intactness of implanted scaffolds. Vascular flowcould not be maintained, however¹⁵. To address this issue,re-endothelialized kidney scaffolds were implanted using the combinationseeding method to maintain blood flow into the implant. This approachdemonstrated scaffold patency for 2 hours after implantation, afterwhich significant clot formation was observed in the parenchyma. Basedon this observation and histological analysis of the kidney scaffolds,it was evident that endothelial cell detachment, which may have occurredfrom the high intravascular pressure and flow, was the primary cause forthrombosis in the renal scaffold. Accordingly, a method was developed tostrengthen the endothelial cell attachment sufficiently to endurephysiologic vascular conditions.

In this study, an antibody conjugation technique was employed to modifythe vascular surfaces of a scaffold in order to enhance endothelial celladhesion. In in vitro testing using a flow chamber system, theconjugation of an endothelial cell-specific antibody against CD31 to thechamber surface prevented endothelial cell detachment under high flowconditions. This in vitro experiment was performed to confirm theeffectiveness of antibody conjugation and the ability of conjugatedantibodies to capture cells in motion. The flow rates used for in vitroflow chamber study are much lower than the cell seeding experimentsperformed with the kidney scaffold. Following implantation ofre-endothelialized CD31 antibody-conjugated kidney scaffolds, prolongedendothelial cell attachment to the antibody-conjugated scaffolds wasobserved that significantly improved vascular patency throughout theparenchyma in vivo. This study shows that the re-endothelializationtechnique disclosed herein, combined with cell-specific antibodyconjugation, results in uniform coverage of endothelial cells within thevasculature of decellularized kidney scaffold and maintains cellattachment leading to prolonged blood flow under physiological vascularconditions. These results have demonstrated that reinforcement ofendothelial cell attachment by antibody conjugation improved vascularpatency following implantation (FIG. 22). However, it is possible that arelative lack of endothelial cell coverage on the conjugated vascularsurface during implantation may attract interaction with other celltypes in the blood, such as platelets, monocytes, and neutrophils. Toreduce the possibility of such interaction, other anti-thromboticsubstances, such as heparin, could be co-conjugated with CD31 antibody.In the platelet adhesion experiment using whole blood that has beendisclosed herein, an antibody-conjugated surface significantly reducedplatelet adhesion when compared to a non-conjugated vascular surface.

The experiments disclosed herein demonstrate the effectiveness ofantibody conjugation-mediated re-endothelialization. One study showedthat the antibody-conjugated re-endothelialized vessels maintainedpatency over a period of 4 hours, while the controls without antibodyconjugation resulted in complete obstruction due to thrombosis. Therenal blood flow (RBF) rate of pig is approximately 470 ml/min¹⁷, andmaintaining renal vascular patency for 4 hours corresponds to 112.8liters of blood flow into the scaffolds. This study further confirms thefeasibility of using porcine-derived decellularized whole kidney as ascaffold for renal tissue engineering. The recellularization andreinforcement methods disclosed herein would serve as an intermediatestep to clinical translation of the decellularized scaffold approach.Previous studies in this area have used small animals, such asrodents³⁻⁵. The porcine kidney transplantable scaffold as disclosedherein provides a renal acellular matrix that has a number ofadvantages, including: (1) similarity to human kidneys in size, (2)lowered infection risk associated with the transplantation process, and(3) heightened relative safety in using pig material, as other acellularporcine tissues, such as heart valve¹⁸ and small intestinal submucosa(SIS) matrix¹⁹, have already been safely used in the clinic. The use ofporcine whole renal scaffolds for renal tissue engineering is reasonablyexpected to translate to an application in humans. We further expectthat an approach termed “semi-xenotransplantation”¹⁶, where autologouscells from the patient could be used to repopulate the acellular porcinekidney matrix, to be safe and effective. The results presented hereindicate that endothelial cell seeding into the vascular network withinacellular porcine kidneys is feasible and results in homogenousformation of endothelium. These results provide evidence that it ispossible to produce a fully endothelialized vascular tree that maintainsvascular patency and delivers nutrients and oxygen to engineeredkidneys, as well as other engineered whole organ systems.

REFERENCES—FOR EXAMPLE 9 ONLY

1. Chazan, J. A. et al., Clin. Nephrol. 35, 78-85 (1991).

2. Ott, H. C. et al., Nat. Med. 14, 213-221 (2008).

3. Petersen, T. H. et al., Science 329, 538-541 (2010).

4. Uygun, B. E. et al., Nat. Med. 16, 814-820 (2010).

5. Baptista, P. M. et al., Hepatology 53, 604-617 (2011).

6. Atala, A. et al., Sci. Transl. Med. 4, 160rv12 (2012).

7. Crapo, P. M. et al., Biomaterials 32, 3233-3243 (2011).

8. Ott, H. C. et al., Nat. Med. 16, 927-933 (2010).

9. Song, J. J. et al., Nat. Med. 19, 646-651 (2013).

10. Ross, E. A. et al., J. Am. Soc. Nephrol. 20, 2338-2347 (2009).

11. Ross, E. A. et al., Organogenesis 8, 49-55 (2012).

12. Nakayama, K. H. et al., Tissue Eng. Part A 16, 2207-2216 (2010).

13. Sullivan, D. C. et al., Biomaterials 33, 7756-7764 (2012).

14. Ko, I. K. et al., Biomaterials 30, 3702-3710 (2009).

15. Orlando, G. et al., Ann. Surg.256, 363-370 (2012).

16. Orlando, G. et al., Transplantation 91, 1310-1317 (2011).

17. Daghini, E. et al., Radiology 243, 405-412 (2007).

18. Lawton, J. S. et al., J. Thorac. Cardiovasc. Surg. 137, 1556-1557(2009).

19. Iannotti, J. P. et al., J. Bone Joint Surg. Am. 88, 1238-1244(2006).

EXAMPLE 10 Whole Organ Engineering—Re-Endothelialization of KidneyScaffolds

Regenerative medicine has shown immense potential to address the limitednumber of transplantable organs and to allow immunosuppression-freetransplantation, through the generation of body parts from patient's ownbiomaterials¹. Among the various approaches to organ bioengineering orregeneration, the seeding of cells on supporting scaffoldingmaterial—namely, cell-on-scaffold seeding technology (CSST)²—seems tooffer the quickest route to clinical application. In fact, thistechnology has allowed the production of numerous, yet relatively simplebody parts for therapeutic purposes, that were eventually implanted inmore than 200 patients¹.

In the wake of these preliminary, yet groundbreaking achievements, CSSTis being applied also to manufacture more complex, metabolic,transplantable organs, including the kidney. In particular,extracellular matrix (ECM) scaffolds obtained through thedetergent-based decellularization of multiple species are used as atemplate for the seeding of kidney-specific cells or progenitor cells,in an attempt to regenerate the parenchymal compartment, as well as ofendothelial cells or progenitor cells aiming at the full regeneration ofthe endothelium. Since the very first report by Ross et al.³ on theproduction of bioactive ECM scaffolds from rodent kidneys, severalstudies have followed⁴⁻¹² and have provided evidence that renal ECMscaffolds can be successfully and consistently produced from virtuallyall species including humans^(9,13), are completely acellular andvirtually non-immunogenic, maintain their architecture and essentialmolecular composition, lack cell membrane molecules, are able todetermine cell phenotype and induce genes of renal development, possessremarkable angiogenic properties as demonstrated by the ability toinduce vessel formation in the chorioallantoic membrane, arebiocompatible in vitro and in vivo, and, when repopulated with renalcells, are able to show some function. Moreover, when acellular porcinerenal ECM scaffolds are implanted in pigs, the framework of the innatevasculature remains well preserved and is able to well sustainphysiologic blood pressure⁴.

In order to validate these promising preliminary data in a moreclinically relevant model, our group is applying CSST to human kidneysinitially procured for transplant purposes, but eventually discarded forvarious reasons¹³. In fact, in the United States, because more than 2600kidneys are discarded annually from the total number of kidneys procuredfor transplantation, we viewed this organ pool as having potential foruse as a platform for renal bioengineering and regeneration research. Weshowed that SDS-based decellularization consistently and successfullyyields human renal ECM scaffolds (hrECMs) with a well-preservedthree-dimensional architecture, an intact glomerular basement membranealong with other important structural proteins. Notably, these scaffoldslack HLA antigens and possess a striking ability to induce angiogenesis,which is an essential biological characteristic of any biomaterials inview of optimal, ad hoc re-cellularization and function followingimplantation in vivo.

Therefore, the disclosed study was conceived, designed and implementedto complete the characterization of hrECMs by evaluating, for the firsttime, the microvasculature using the resin casting method, in additionto robustly assessing the dimension of the glomerular capillaries andarterioles. Moreover, in order to thoroughly assess the compliance andresilience of the framework of the innate vasculature within hrECMs, theactual arterial and venous pressures were measured within the frameworkof the vasculature of our matrices, with the use of a pulse-wave, setwithin the physiological limits. Finally, an assessment was made of thepresence of soluble growth factors (GFs) that possibly can be retainedwithin the rhECMs and that are responsible for inducing angiogenesis.

Kidney Procurement and Preparation

Kidneys were procured for transplant purposes but then discarded forvarious reasons including anatomical abnormalities such asglomerulosclerosis, interstitial fibrosis, tissue inflammation orcortical necrosis. All organs were procured within the designatedservice area of our local procurement organization (Carolina DonorService) and were refused by all local, regional, and nationaltransplant centers. Kidneys were offered to the transplant team of theWake Forest School of Medicine and processed at the Wake ForestInstitute for Regenerative Medicine after the complete exhaustion of thenational list.

Kidneys were received in sterile cold solution (saline solution NaCl0.9%) and preserved until shipment. The aortic patch and the renal veinwere prepared according to transplant protocol. The renal vein wasdissected and sectioned at 2 cm from its origin. Multiple arteries werereconstructed in order to create a single arterial inlet. Peripheral fatand lymphatic tissue were ligated with 2/0 silk ties. Sixteen gaugeintravenous catheters were inserted into the renal artery, the renalvein, and the ureter. The renal artery and the ureter were subsequentlytested for possible leakages and eventually repaired with 6/0 Prolenesutures. Because all kidneys had been biopsied at the upper pole at thetime of procurement, a renorrhaphy of the “wedge” defect was performedwith 4-0 PDS suture in a running way. Kidneys were finally placed on iceuntil decellularization.

Kidney Decellularization and hrECMs Production

The angiocatheters previously inserted in the renal artery and in theureter were connected to a pump (Masterflex L/S peristaltic pump withMasterflex L/S easy load pump head and L/S 16G tubing, Cole-PalmerInstrument Co, Vernon Hills, Ill., USA) to allow continuous rinsing withdifferent solutions, starting with phosphate buffer saline (PBS) at therate of 12 ml/minute for 12 hours (8,640 mL total).

Afterward, 0.5% sodium dodecyl sulfate (SDS, Sigma-Aldrich, St. Louis,Mo., USA)-based solution was delivered at the same flow rate for 48hours (34,560 mL total) in both the renal artery and ureter. Finally thekidneys were rinsed with DNase (Sigma-Aldrich, St. Louis, Mo., USA) for6 hours at a flow rate of 6 ml/hour and then with phosphate buffersaline, PBS (Sigma-Aldrich, St. Louis, Mo., USA) at the same flow ratefor 5 days (43,320 mL total). The histological characterization of thebioscaffolds was performed as previously described¹³ in order to confirmabsence of cellular residual.

Resin Vascular Cast

Resin casting of the innate vasculature was obtained as previouslydescribed^(14, 15). A total of 16 kidneys underwent resin vascular casttreatment divided into two different groups: group 1 counting eightnative/cellular kidneys versus group 2 counting eight hrECMs. Precastingtreatment was carried out via injection through arterial inlet of 60 mlheparinized PBS solution in order to prevent any kind of blood clottingwhile washing out the remaining blood from the vasculature tree. Foreach kidney, 60 ml of specific casting resin was prepared by mixing 50ml of casting resin monomer (Batson's #17 Monomer Base SolutionCat#02599, Polyscience, Inc., Warrington, Pa., USA), 10 ml of catalyst(Batson's #17 Anatomical Corrosion Kit Promoter Cat# 02610, Polyscience,Inc., Warrington, Pa., USA) and 10 drops of promoter (Batson's #17Catalyst Cat# 02608, Polyscience, Inc., Warrington, Pa., USA). Thissolution was then slowly injected through the renal artery (15-20 mlcolored by red dye, Batson's #17 Anatomical Corrosion kit Red dye Lot#533945, Polyscience, Inc., Warrington, Pa., USA) the renal vein (15-20ml colored by blue dye, Batson's #17 Anatomical Corrosion kit Blue dyeLot #623514, Polyscience, Inc., Warrington, Pa., USA) and the ureter(10-15 ml colored by yellow dye, Batson's #17 Anatomical Corrosion kitYellow dye Lot #623514, Polyscience, Inc., Warrington, Pa., USA). Afterthe onset of polymerization kidneys were placed in deionized waterovernight and then all the tissue was cleared by two alternating rinses(24 hours each) with 10% and 5% of hydrochloric acid, respectively.

Scanning Electron Microscopy (SEM)

Five 5×5×5 mm samples were obtained from each kidney cast (a total ofeighty samples, forty from group 1 and forty from group 2) in randomlyselected areas of the cortex simply cutting them out with microsurgicalscissors. These samples were mounted on silver plates sputter coatedwith gold and analyzed by Scanning Electron Microscopy (SEM) at 15 KV(Hitachi S-2600N Scanning Electron Microscopy, Hitachi, Chiyoda, Tokyo,Japan). From each sample, two three-dimensional images of the afferentartery and the glomeruli were taken (for a total of 160 glomerularimages captured). All of the images were then analyzed by Image-Jsoftware (http://rsb.info.nih.gov/ij/). Afferent artery diameter wasmeasured at three different points and averaged for statisticalanalysis. Glomerular diameter measured in the long and short axis andsubsequently averaged. Modeling each glomerulus as a sphere, thesediameter averages were used to calculate volume ( 4/3 πr3). Six randomlyselected glomerular capillaries from each image were measured and usedto calculate average capillary diameter. From these data, themorphological properties of the glomeruli and their afferent arterieswere evaluated.

Determination Of Machine-Perfusion Vascular Responsiveness

Nine donated human kidneys were machine-perfused at 4° C. for 12 hourswith modified Krebs solution, using a unique cardioemulation perfusiontechnology (VasoWave®, Smart Perfusion, Denver, N.C.). This systemproduces a cardioemulating pulse wave to generate physiological systolicand diastolic pressures and flow rates within the organ. The system iscapable of controlling the oxygen content of the perfusate above andbelow physiological norms. During perfusion, arterial pressuremeasurements were taken. A comparison was made between machine-setpressures (systolic/diastolic) and actual pressures within organs underperfusion. Organs with intact vasculature replicate an elastic responseto machine-set pressures. Following decellularization, scaffolds fromthe same set of nine human kidneys were again machine-perfused (modifiedKrebs solution at 4° C., for 12 hours) and arterial pressuremeasurements were taken as described. It is important to note that, ifscaffolds had been damaged by decellularization, they would notelastically respond to machine-set pressures, and would leak fluid andwould fail to sustain pressurization. Pressure measurements werecollected for both cellularized and decellularized kidneys at a rate of100/second and then sampled every 1000 points to create mean summarydata.

Growth Factor Retention

5 hrECMs and 5 samples of native kidneys before decellularization wereused to evaluate growth factor retention. Samples were procured with a 7mm biopsy punch, stored in sterile PBS with 2% pen/strep (HyclonePe/Strep solution, Fisher Scientific, Waltham, Mass., USA) and thenshipped to the manufacturer (Raybiotech, http://www.raybiotech.com/) andprocessed accordingly. Specifically, tissue biopsies underwenthomogenization by sonication in RayBiotech's proprietary lysis buffer(500 μL of lysis buffer per 10 mg tissue) and centrifugation for 5minutes at 10,000×g. Supernatants were then collected and assayedimmediately or frozen for future use. Protein expression profiles werecollected using the RayBio® Human Growth Factors Antibody Array G series1, a custom glass chip-based multiplex ELISA array which measures 40cytokines simultaneously. The tissue supernatants were incubated withthe array chips after a blocking step and these were washed to removenonspecific proteins, and biotin-labeled detection antibodies wereadded. The cytokine-antibody-biotin complexes can then be visualizedthrough the addition of a HiLyte Fluor 532™ dye-labeled streptavidin.Spot intensities extracted from the scanned array image were normalizedto positive controls included within each array. Average fluorescentintensity was obtained from duplicate signal intensities, adjusted toremove background and normalized to a positive control to account fordifferences among sub-arrays.

Immunofluorescence to Assess Maintenance of Structural Specific hrECMsComponents

In order to evaluate the presence of specific hrECM proteins, 1 cm³biopsies taken from the cortex of de-cellularized human kidneys werefixed in 10% formalin (Azer scientific, PA) for 2 hours. After fixation,samples were dehydrated in alcohol gradients and placed in toluene(Sigma-Aldrich, St. Louis, Mo.) for 30 minutes. Samples were keptovernight in a 50:50 toluene:paraffin mixture. Samples were next changedinto paraffin for 2 hours and later embedded. 5 μm thick sections(Rotary Microtome-Leica, Rotary Microtome RM2235) were deparaffinizedand rehydrated in alcoholic gradients for histology. Immunofluorescenceanalysis was conducted by overnight incubation with primary antibodiesCollagen IV (ABCAM, Cambridge, Ma. 1:100), VEGFR-2 (ABCAM, Cambridge,Ma. 1.5:100), VE-Cadherin (ABCAM, Cambridge, Ma. 1.5:100) followed by 30minute incubation with secondary anti-mouse or anti-rabbit Alexa Fluor555 (Life Technologies Grand Island, NY 1:500). DAPI mounting (VectorLaboratories Burlingame, CA) was used to visualize samples with a LeicaDM5500 B Microscope System.

Statistics

All graphical data are displayed as the mean+SEM. Statistical analysiswas performed using the Student's T-test and MatLab Software in order tocompare measurements of glomeruli, arterioles and capillaries betweenthe two experimental groups as well as the machine-perfusion vascularresponsiveness. A p value less than 0.05 was considered statisticallysignificant.

Cast Preparation and Morphometric Analysis

The resin filled the framework of the innate vasculature throughout thewhole parenchyma, ultimately producing a high-fidelity three-dimensionalcasting, as shown in FIGS. 40A-D. The corrosion cast protocolsuccessfully produced 16 satisfactory whole-kidney casts (8 nativekidneys and 8 scaffolds), with uniform representation of the vascularnetwork and glomeruli through the entire cortex, as shown in FIGS.40E-H.

Five biopsies were taken from each organ and scaffold. Of each biopsy,five SEM images of randomly selected glomeruli were captured and studied(FIG. 28). Morphometrical endpoints were: sagittal and transversalglomerular diameter (green lines); diameter of the afferent artery (redlines); diameter of six different glomerular capillaries, randomlyselected (yellow lines). Results are shown synoptically in FIG. 29. Theaverage afferent arteriolar diameter was 24.20±0.49 μm in native kidneysversus 23.65±0.63 μm in hrECMs (p=n.s.). The average glomerular diameterwas 224.37±5.23 μm in the native kidneys versus 182.93±3.8 μm in thescaffolds (p<0.01). Volumetric calculations were carried out using thesefigures by modeling glomeruli as spheres. Analysis showed that volume ofnative glomeruli was statistically higher that in the hrECMs(7.17×10⁶±6.62×10⁵ μm³ versus 3.81×10⁶±3.07×10⁵ μm³, p<0.01). Meancapillary width was 11.36±0.20 μm for native kidneys and 11.37±0.20 μmfor renal scaffolds (p=n.s.). The mean afferent arteriolar diameter was24.20±0.64 μm for native kidneys and 23.65±0.72 μm for renal scaffolds(p=n.s.).

Machine-Perfusion Vascular Responsiveness

In order to evaluate the effects of decellularization, the vascularelasticity and ability to sustain pressure in both intact(pre-de-cellularization) and processed (post-de-cellularization) kidneyswas measured. To accomplish this, the VasoWave perfusion system wasconnected to the arterial and venous vessels and pulsing was appliedwhile measuring pressure responsiveness and fluid load in theclosed-loop system. Machine-perfused kidneys and hrECMs stabilizedwithin 20 minutes of anastomosis and were effectively perfused in aclosed-circuit system using 1-1.5 L of modified Krebs solution for 12hours. No interstitial edema or swelling was noted and there was nofluid ‘weeping’ from the surfaces or under the capsule of theexperimental groups.

As shown in FIGS. 30A and 30B, the vasculature of both native kidney andhrECM demonstrated an elastic response to machine-set pressures. With amachine-set systolic pressure of 90 mmHg, the mean vascular elasticresponse of the native organ (measured by the difference of set versusactual pressure) was 0.898%. The mean vascular elastic response of thenative organ to the diastolic (50 mm Hg) pressure wave was 7.47%. Ofnote, administration of the arterial pressure wave temporally resultedin slightly higher actual arterial pressures, due to arterial dilationopposed by tissue pressure and venous backpressure. This is commonlyseen as a manifestation of vascular resistance in normal kidneys.

Following de-cellularization, vascular elastic responses were againevaluated in the same 9 kidneys. With a machine-set systolic pressure of90 mm Hg, the mean vascular elastic response of the hrECMs (measured bythe difference of set versus actual pressure) was 1.76%. There was notemporal rise in actual arterial pressure, demonstrating thatdecellularization effectively removed tissue backpressure. The meanvascular elastic response of the hrECMs to the diastolic (50 mm Hg)pressure wave was 2.48%. Removal of cellular material did not compromisethe elastic response of the vascular scaffold. In fact, smoothing of theelastic response was indicative of more effective perfusion (lessdifference in the diastolic mean deviation from the machine-set pressurewave).

Growth Factor Analysis

The multiplex array shows hrECMs retain numerous GFs that play vitalrole during important biological processes, including angiogenesis,renal development and regeneration, as well as glucose homeostasis (FIG.31 and Table 2). Immunofluorescence confirmed the presence of collagenIV, VEGF-R2, as well as Ve-Cad (FIGS. 31B-D) within the hrECMs,indicating the preservation of the glomerular basement membrane and thepresence of key molecules that support the capillaries structure withinthe glomeruli.

The present study was conceived and designed to address critical, yetunaddressed aspects of ECM characterization, namely the integrity andresilience of the innate vasculature through vessel morphometry andperfusion studies, and the ability of hrECMs to retain GFs throughdirect quantification of a custom-made panel of relevant GFs and ad hoc“in tissue” staining. The results show that the framework of the innatevasculature of hrECMs is well-preserved and retains its innateresilience, and that GFs that are key players in critical processes oftissue development like angiogenesis remain within the matrixpost-de-cellularization at significant concentrations.

TABLE 2 Human Growth Factors Array - Native kidney AR bFGF b-NGF EGF EGFR FGF-4 FGF-6 438.10 ± 16.25  5585.10 ± 1363.04 462.70 ± 29.95  1576.10± 326.27  5360.30 ± 577.50  565.60 ± 20.19  54.10 ± 9.80  FGF-7 GCSFGDNF GM-CSF HB-EGF HGF IGFBP-1 389.70 ± 11.84  499.80 ± 23.68  569.30 ±29.04  402.90 ± 22.49  73.90 ± 10.36 16626.40 ± 3402.84  10023.00 ±5834.07  IGFBP-2 IGFBP-3 IGFBP-4 IGFBP-6 IGF-I IGF-I SR IGF-II 2018.9 ±555.18 21.3 ± 5.72 266.1 ± 12.27 266.5 ± 20.72  346 ± 40.67 497.9 ±8.28  63180.1 ± 1964.59 M-CSF M-CSF R NT-3 NT-4 PDGF R a PDGF R bPDGF-AA  411.7 ± 151.93  523.1 ± 169.12 252.9 ± 11.33 469.3 ± 11.83164.5 ± 12.02 433.7 ± 21.52 554.6 ± 48.24 PDGF-AB PDGF-BB PIGF SCF SCF RTGF-a TGF-b  984.2 ± 138.75  641.5 ± 170.89  281 ± 14.87 496.4 ± 68.47386.5 ± 24.35  44.9 ± 11.85 4520.8 ± 391.55 TGF-b 2 TGF-b 3 VEGF VEGF R2VEGF R3 VEGF-D 434.9 ± 10.63 125.3 ± 10.99 28.9 ± 9.62  855.4 ± 107.12 317 ± 29.87 185.8 ± 24.84 Human Growth Factors Array - Acellular kidney(hrECM) AR bFGF b-NGF EGF EGF R FGF-4 FGF-6 753.6 ± 29.40 612.6 ± 50.56654.3 ± 40.35  948.5 ± 116.63 255.8 ± 37.50 1051.8 ± 140.84 277.2 ±60.52 FGF-7 GCSF GDNF GM-CSF HB-EGF HGF IGFBP-1 500.2 ± 22.03 732.2 ±34.15 859.1 ± 35.39  503 ± 57.59 233.3 ± 36.17 802.6 ± 88.85 791.5 ±78.59 IGFBP-2 IGFBP-3 IGFBP-4 IGFBP-6 IGF-I IGF-I SR IGF-II 582.9 ±98.18 106.7 ± 16.00 502.9 ± 44.40  905.7 ± 184.62 483.5 ± 47.51  958.4 ±131.08 17775.2 ± 4617.7  M-CSF M-CSF R NT-3 NT-4 PDGF R a PDGF R bPDGF-AA 235.9 ± 18.84 445.2 ± 47.80  785.2 ± 121.75 713.2 ± 29.28  80.3± 13.56 475.4 ± 47.61  746.8 ± 108.95 PDGF-AB PDGF-BB PIGF SCF SCF RTGF-a TGF-b 957.9 ± 97.99  490 ± 67.16  904.6 ± 148.18 876.3 ± 67.80 929.6 ± 110.23 268.6 ± 39.50 8322.7 ± 297.22 TGF-b 2 TGF-b 3 VEGF VEGFR2 VEGF R3 VEGF-D 611.8 ± 16.95 351.1 ± 32.06  78.8 ± 17.97 542.2 ±51.91  586 ± 76.75  538.5 ± 150.54

In regenerative medicine, CSST has shown the greatest potential forclinical translation, allowing the production of body partsbioengineered from patient's cells that were eventually implanted inmore than 200 patients without any anastomosis to the recipient'svasculature, at the time of implantation. In contrast, more complexmetabolic organs like the kidney do require re-anastomosis between thevascular pedicle of the bio-artificial organ and the recipient'sbloodstream, to allow function. Therefore, the acellular framework ofthe innate vasculature endowed within ECM scaffolds used for tissueengineering purposes should maintain the basic characteristics andresilience of the intact counterpart, in order to allow implantationwith physiological blood pressure being sustained in vivo. Moreover, asone of the critical functions of ECM in mammals is to act as reservoirfor GFs to be released following specific stimuli¹⁶, ECM scaffolds usedfor tissue engineering should maintain GFs. As hrECMs were proven tohave the ability to induce the formation of new vessels in the in ovo¹³bioassay, there is evidence that several molecules involved in theinflammatory and angiogenesic cascades are present in the matrix postdecellularization, demonstrating that the disclosed matrices arebioactive and so may facilitate the regeneration and re-building of newtissue after re-cellularization and ultimately implantation.

A morphometric study of the framework of the innate vasculature wasconducted by three-dimensional analysis of SEM images of corrosioncasting, as described by Pereira-Sampaio et al. and Manelli et al.Analysis and measurement of casting imaging at SEM revealed thatmorphology and dimensions of the acellular glomerulus and its vascularnetwork are relatively well-preserved post-de-cellularization and arecomparable to their cellular counterparts. The diameter and volume ofthe glomerular macrovasculature within the hrECMs were found to becontracted when compared with the normal kidney. This finding may be dueto the absence of cells, accounting for a significant part of the wholevolume of the intact, cellular glomerulus. No significant differences inthe width of glomerular capillaries and the branching pattern andintegrity of the larger vessels was detected. Overall, these dataconfirm the operability of the de-cellularization method yielding hrECMsthat provide a framework of the vascular network preserved at allhierarchical levels, namely large vessel and small cortical vessels. Thepreservation of an intact vascular bed in the disclosed hrECMs isimportant because use of the innate scaffolds in vivo depends on anintact vascular network that is able to sustain physiological bloodpressure and perfusion. With regard to the decreased glomerular diameterand volume observed in the disclosed matrices, it is reasonable toexpect that the regeneration of the endothelium would normalize thosevalues (i.e., bring them to values observed in the innate, cellularkidney).

As the kidney is one of the major control stations of intravascularblood pressure, experiments were designed to determine whether theintact framework of the vascular network of hrECMs maintains functionand resilience, namely its ability to respond to changes inintravascular pressure. To do so, a state of the art technology was usedthat provided both a cardioemulating, physiologic pulse pressure and theability to dynamically measure applied pressure and elastic vascularresistance for each pulse wave. Our analysis of pulse wave data andresponse to pressurization demonstrated that, although the process ofde-cellularization removed cells, the scaffolds demonstrated the sameelastic response characteristic of intact vascular beds. Specifically,scaffolds did not leak (no fluid wept from the surface) during perfusionand elastic rebound in response to an applied pulse wave was seen,although slightly diminished when compared to intact, cellularizedkidneys.

To finalize the characterization of the matrices and determine whetherthey have the characteristic of implantable organs and induce formationof new vessels, the GFs content of hrECMs was examined. It is importantto emphasize that, during kidney development as well as in the naturalhistory of chronic kidney diseases, important GFs secreted and retainedwithin the ECM orchestrate very complex cell-cell and cell-matrixinteractions. For example, variation in concentration of several growthfactors, including transforming growth factor-α (TGF-α), heparin-bindingepidermal growth factor (HB-EGF), insulin-like growth factor (IGF), andfibroblast growth factor (FGF), are responsible for cell migration,proliferation, differentiation, induction of pro-fibrotic processes aswell as possible pro-healing signaling with scar resolution¹⁷.Therefore, as GFs play a major role in determining progression orblockade of kidney damage, it is of vital importance to determine if thede-cellularization process of human kidneys preserves important stimulithat could eventually facilitate cellular repopulation and mediateinduction of fibrosis, in future in vivo applications. Interestingly,several GFs, including TGF-α, FGF-6, insulin-like growth factor bindingprotein 3 (IGFBP-3), HB-EGF, insulin-like growth factor binding protein6 (IGFBP-6), neurotrophin 3 (NT-3), placenta growth factor (P1GF),transforming growth factor-β (TGF-β), vascular endothelial growth factor(VEGF), and vascular endothelial growth factor-D (VEGF-D), were observedto be present within the hrECMs. The presence of critical transmembraneglycoproteins was confirmed, i.e., Ve-cadherin and VEGFR-2, which arevital for the function and the strength of the endothelium and for themaintenance of glomerular endothelial cells fenestrations, essential forglomerular barrier filtration. For example, VEGFR-2 postnatal deletiondemonstrates a global defect in the glomerular microvasculature and theparacrine VEGF-VEGFR-2 signaling loop is identified as a criticalcomponent in the developing and in the filtering glomerulus¹⁸. Thepresence of these GFs should not be underestimated, showing that hrECMsprovide not only structurally supportive vasculature, but also maintainarchitecturally specific transmembrane glycoproteins for glomerularendothelial cell function and support. Nevertheless, these data reveal,for the first time, that hrECMs are a candidate for tissue engineeringpurposes and possess all the necessary characteristics to inducefunctional vasculature in vivo.

The disclosed studies demonstrate that the framework of the innatevasculature of hrECMs is maintained at all hierarchical levels, isresilient, and can sustain intravascular pressures comparable to what isobserved in normal physiology. Also, hrECMs retain numerous GFs that arenecessary for the maintenance of endothelial cell homeostasis andfunction. These results establish that the scaffolds prepared asdisclosed herein are useful in the regeneration of the cellularcompartment and in vivo implantation of bioengineered renal organoidsand organs for transplant purposes.

REFERENCES—FOR EXAMPLE 10 ONLY

-   -   1. Orlando G, Soker S, Stratta R J. Organ bioengineering and        regeneration as the new Holy Grail of organ transplantation Ann        Surg. 2013 August; 258(2):221-32.    -   2. Salvatori M, Peloso A, Katari R, Zambon J P, Soker S, Stratta        R J, Orlando G. Semixenotransplantation: the regenerative        medicine based-approach to immunosuppression-free        transplantation and to meet the organ demand.        Xenotransplantation, Jul. 8, 2014 ahead of print.    -   3. Ross E A, Williams M J, Hamazaki T, Terada N, Clapp W L, Adin        C, et al. Embryonic stem cells proliferate and differentiate        when seeded into kidney scaffolds. J Am Soc Nephrol 2009;        20:2338-47.    -   4. Orlando G, Farney A, Sullivan D C, AbouShwareb T, Iskandar S,        Wood K J, et al. Production and implantation of renal        extracellular matrix scaffolds from porcine kidneys as a        platform for renal bioengineering investigations. Ann Surg 2012;        256:363-70.    -   5. Nakayama K H, Batchelder C A, Lee C I, Tarantal A F.        Decellularized rhesus monkey kidney as a three-dimensional        scaffold for renal tissue engineering. Tissue Eng Part A 2010;        16(7):2207-16.    -   6. Nakayama K H, Batchelder C A, Lee C I, Tarantal A F. Renal        tissue engineering with decellularized rhesus monkey kidneys:        age-related differences. Tissue Eng Part A 2011;        17(23-24):2891-901.    -   7. Nakayama K H, Lee C C, Batchelder C A, Tarantal A F. Tissue        specificity of decellularized rhesus monkey kidney and lung        scaffolds. PLoS One 2013; 8(5):e64134.    -   8. Sullivan D C, Mirmalek-Sani S H, Deegan D B, Baptista P M,        Aboushwareb T, Atala A, Yoo J J. Decellularization methods of        porcine kidneys for whole organ engineering using a        highthroughput system. Biomaterials 2012; 33(31):7756-64.    -   9. Song J J, Guyette J P, Gilpin S E, Gonzalez G, Vacanti J P,        Ott H C. Regeneration and experimental orthotopic        transplantation of a bioengineered kidney. Nat Med 2013;        19(5):646-51.    -   10. Bonandrini B, Figliuzzi M, Papadimou E, Morigi M, Perico N,        Casiraghi F, Dipl C, Sangalli F, Conti S, Benigni A, Remuzzi A,        Remuzzi G. Recellularization of well-preserved acellular kidney        scaffold using embryonic stem cells. Tissue Eng Part A 2014;        20(9-10):1486-98.    -   11. Wang Y, Bao J, Wu Q, Zhou Y, Li Y, Wu X, Shi Y, Li L, Bu H.        Method for perfusion decellularization of porcine whole liver        and kidney for use as a scaffold for clinical-scale        bioengineering engrafts. Xenotransplantation. Oct. 7, 2014 [Epub        ahead of print].    -   12. Choi S H, Chun S Y, Chae S Y, Kim J R, Oh S H, Chung S K,        Lee J H, Song P H, Choi G S, Kim T H, Kwon T G. Development of a        porcine renal extracellular matrix scaffold as a platform for        kidney regeneration. J Biomed Mater Res A. Jul. 16, 2014 [Epub        ahead of print].    -   13. Orlando G, Booth C L, Wang Z, Totonelli G, Ross C L, Moran        E, Salvatori M, Maghsoudlou P, Turmaine M, Delario G,        Al-Shraideh Y, Farooq U, Farney A C, Rogers J, Iskandar S S,        Burns A, Marini F C, De Coppi P, Stratta R J, Soker S. Discarded        human kidneys as a source of ECM scaffolds for kidney        regeneration technologies. Biomaterials 2013; 34:5915-25.    -   14. Pereira-sampaio M A, Henry R W, Favorito L A, Sampaio F J.        Cranial pole nephrectomy in the pig model: anatomic analysis of        arterial injuries in tridimensional endocasts. J Endourol. 2012;        26(6):716-21.    -   15. Manelli A, Sangiorgi S, Binaghi E, Raspanti M. 3D analysis        of SEM images of corrosion casting using adaptive stereo        matching. Microsc Res Tech. 2007; 70(4):350-4.    -   16. Hynes R O. The extracellular matrix: not just pretty        fibrils. Science. Nov. 27, 2009; 326(5957):1216-9.    -   17. Jain R K, Au P, Tam J, et al. Engineering vascularized        tissue. Nat Biotechnol. 2005; 23(7):821-3.    -   18. Rahimi N, Kazlauskas A. A role for cadherin-5 in regulation        of vascular endothelial growth factor receptor 2 activity in        endothelial cells. Mol Biol Cell. 1999; 10:3401-3407.

EXAMPLE 11 Whole Organ Engineering—Re-Endothelialization of LiverScaffolds

Donor shortage remains a continued challenge in liver transplantation.Recent advances in tissue engineering have provided the possibility ofcreating functional liver tissues as an alternative to donor organtransplantation. Small bioengineered liver constructs have beendeveloped, however a major challenge in achieving functionalbioengineered liver in vivo is the establishment of a functionalvasculature within the scaffolds. Our overall goal is to bioengineerintact livers, suitable for transplantation, using acellular porcineliver scaffolds. We developed an effective method for reestablishing thevascular network within decellularized liver scaffolds by conjugatinganti-endothelial cell antibodies to maximize coverage of the vesselwalls with endothelial cells. This procedure resulted in uniformendothelial attachment throughout the liver vasculature extending to thecapillary bed of the liver scaffold and greatly reduced plateletadhesion upon blood perfusion in vitro. The re-endothelialized livers,when transplanted to recipient pigs, were able to withstandphysiological blood flow and maintained for up to 24 hours. This studydemonstrates, for the first time, that vascularized bioengineeredlivers, of clinically relevant size, can be transplanted and maintainedin vivo, and represents the first step towards generating engineeredlivers for transplantation to patients with end-stage liver failure.

Liver transplantation represents the only curative treatment forend-stage liver disease. Unfortunately this solution is limited by acritical shortage of donor organs that are suitable for transplant.According to the United Network for Organ Sharing, over 16,000 patientsare currently awaiting liver transplant, while less than 7000 donororgans become available annually. This discrepancy between organ supplyand demand results in thousands of deaths each year. Several treatmentstrategies are being developed to sustain critically ill patients untila time when a transplantable donor organ is available [1, 2]. However,these therapies can only buy a small amount of additional time forpatients in liver failure. Recently, the use of non-heart-beating liverdonors (NHBD) has been considered, but this would still not close theorgan supply/demand gap [3-6]. Clearly, alternative treatments forpatients with end stage liver disease need to be investigated. Over thepast decade, the fields of regenerative medicine and tissue engineeringhave offered new strategies for the generation of engineered organs[7-9]. These new strategies are based upon the use of scaffolds with apreexisting architectural structure that are seeded with an appropriatepopulation of cells [10]. Natural tissue extracellular matrices (ECM)possess a dynamic network of macromolecules with organ-specificanatomical and biochemical properties [11, 12]. It would be advantageousto include these properties in any scaffold considered for organengineering.

Whole organ engineering represents the ultimate solution for completelysolving the shortage of transplantable organs [13-20]. For the reasonsmentioned above, decellularized whole organ matrices would be thepreferred option for a construct scaffold. These scaffolds are easilygenerated by perfusion of donor organs with mild detergents that removethe cellular components from the organ [21, 22]. Importantly,decellularized whole organ matrices retain the vascular network andtissue microarchitecture present in the native organ [23, 24]. Thecurrent study is based on the development of a decellularized liverscaffold that is subsequently repopulated with cells isolated from livertissue samples. The ultimate goal of these studies is to create atransplantable organ that could replace the functional capabilities ofthe patient's failing liver [25, 26].

The decellularization process for the production of an acellular organscaffold has been under development for many years, and the results havebeen quite remarkable [20, 21]. However, the recellularization processhas proven to be very challenging. One critical obstacle to achieving atransplantable, recellularized organ is reestablishment of a patentvasculature with sufficient endothelial cell coverage to preventthrombosis. Acellular ECM is potently thrombogenic, and blood clots willform in an insufficiently endothelialized construct, even with the useof standard anticoagulant therapy [27].

The goal of the current study was to re-establish a functionalvasculature in bioengineered livers of clinically relevant size. Toachieve this goal, we developed a novel re-endothelialization techniquebased on using anti endothelial cell antibodies to stabilize seededcells on the vessel walls. Vascular functionality was validated bytransplanting the re-endothelialized livers using a heterotopictransplantation model with inflow from the renal artery and the outflowto the renal vein (FIG. 32). The major findings indicated that there-endothelialized liver scaffolds were able to withstand physiologicblood pressure and maintain blood flow within the bioengineered liversfor 24 hours. This study shows, for the first time, a strategy forovercoming a major hurdle in the engineering of transplantable liver,the establishment of functional vasculature.

Decellularization of Porcine Liver

Native livers were harvested from 5 to 8 kg piglets. The portal vein(PV), common hepatic artery (HA), suprahepatic (SH)— and infrahepatic(IH) inferior vena cava (IVC) were cannulated with smart site connectors(Cole Parmer) attached to 14 G tubing with inflow and outflow adjustedto mimic normal flow through the organ. Detergent solutions (1% TritonX-100 and 0.1% ammonium hydroxide in distilled water) were perfused intothe liver tissue using a peristaltic pump (Master flex L/S with Masterflex L/S easy load pump head, Cole Parmer, Vernon Hills, Ill., USA).Decellularization of the liver was performed by perfusing the organ at aflow rate of 0.5 ml/min for 2-3 days, followed by washing with salinefor 3-4 days. The decellarized livers were sterilized by gammairradiation at 1.2 MRad (12,000 Gy) prior to cell seeding.

Characterization of Acellular Liver Scaffold

To evaluate the efficiency of scaffold decellularization, DNAquantification and histological analysis (H&E) were performed. For DNAquantification, samples were excised from representative lobes of nativeand decellularized livers. The samples were minced and lyophilized inpreparation for analysis (Labconco, Kansas City, Mo.). DNA was extractedfrom 5 mg samples using the Qiagen DNeasy Blood and Tissue Kit (QiagenInc., Valencia, Calif.) and quantified using Quant-iT PicoGreen(Invitrogen Corp., Carlsbad, Calif.). Fluorescence from the PicoGreensignal indicates residual DNA and was measured at 525 nm (excitation 490nm) using a SpectraMax M5 Multi-Mode Microplate Reader (MolecularDevices Inc., Sunnyvale, Calif., US).

To evaluate the maintenance of vasculatures within the decellularizedliver scaffold, angiographic studies were performed using computedtomography (CT) and liver vascular casting combined with electronmicroscopic analysis. For CT imaging of native and decellularized liverscaffold, a CT contrast agent (MICROFIL, Flowtech, Inc., Carver Mass.)was infused through the PV at 1 ml/min flow rate, while the artery andveins were clamped. The livers were scanned on a Toshiba Aquilion 32 CTscanner (Toshiba America Medical Systems, Inc., Tustin, Calif.) andanalyzed with CT angiography body and soft tissue deformation processingalgorithms. Imaging was conducted using a TeraRecon Aquarius Workstation(TeraRecon Inc., Foster City, Calif.).

Liver casts were prepared using Batson's compound no 17 (PolysciencesInc., Warrington, Pa.) injected through the portal vein with constantflow rate (1 ml/min) using a syringe pump (New Era Pump Systems, Inc.,Farmingdale, N.Y., USA). Following the resin injection, the infusedliver was incubated to allow the polymerization within the livervascular system. After curing, the liver tissue was digested with 20%sodium hydroxide at 37° C., followed by washing with deionized water.The vasculatures of the liver cast were imaged using scanning electronicmicroscopy (SEM). The cast segments were mounted on silver plates,sputter coated with gold and imaged at 15 KV.

Re-Endothelialization of Acellular Liver Scaffold

To improve re-endothelialization of vasculatures within the liverscaffold, rat anti-mouse CD31 antibody (BD sciences, Franklin Lakes,N.J.) was conjugated to the acellular liver scaffold. Liver scaffoldswere treated with a solution of1-Ethyl-3-[3-dimethylaminopropyl]carbodiimide hydrochloride (EDC)(SigmaeAldrich) and Nhydroxysuccinimide esters (NHS) (SigmaeAldrich) inphosphate buffered solution (PBS) for 30 minutes. Subsequently, 20 ml ofa 50 mg/mL antibody solution was injected through the artery and veins,followed by incubation for 2 hours at room temperature. Constructs werewashed by perfusion with PBS to remove unreacted antibodies and smallpieces of the antibody-conjugated constructs were embedded into OCTcompound for preparation of frozen blocks, followed by sectioning of thefrozen blocks. CD31 antibody conjugation on the vasculatures within thesections was confirmed by treatment with Alexa588 goat anti-rat antibody(BD Biosciences). The conjugated antibody was semi-quantified by imagingthree randomly selected areas (n=3 livers per group) with a fluorescencemicroscope (Leica). Fluorescence intensity was determined using Image Jsoftware. For re-endothelialization of the antibody-conjugated liverscaffold, vascular endothelial cells expressing GFP protein (MS1) wereused. MS1 seeding was performed using a two-step process: a) static andb) perfusion (FIG. 32). For static cell seeding, 7, 3, 5, and 5 ml of10×10⁶/ml MS1 cells suspended in DMEM-high glucose medium (InvitrogenLife Technologies, Carlsbad, Calif.) supplemented with 10% FBS(Invitrogen Life Technologies) were infused into PV, HA, SH-IVC, andIH-IVC, respectively. The seeded cells were then allowed to attach for 1hour, followed by another 1-hour infusion performed following a180-degree rotation of the liver scaffold from the original position.This allows for a more even distribution of cells within the construct.Following the static seeding, the liver scaffold was connected to abioreactor system and equilibrated in 10% FBS DMEM medium at 1 ml/minovernight. Perfusion bioreactors consisted of a glass chamber for theliver, with media circulation via platinum-cured silicone tubing(Cole-Parmer, L/S 15) using Cole-Parmer Digital Drive peristaltic pumps.Bioreactors contained 1.5 L of growth medium and were maintained inhumidified conditions at 37° C., supplemented with 5% CO₂. Media wasperfused directly into the liver via the PV under constant stirring viaa magnetic stirrer. For perfusion cell seeding, 5 ml of a suspension of10×10⁶/m1 cell perfused through the PV at 3 ml/minute. The flow rate wasgradually increased to 5 and 10 ml/minute at 10-12 hour intervals, andmaintained at 10 ml/minute for 3 days. After the perfusion culture,several MS1 seeded liver scaffolds were fixed in 10% formalin, embeddedin OCT compound, and processed for histological analysis (H&E and DAPInuclear staining). Efficiency of re-endothelialization of the acellularliver scaffold was determined by histological analysis [H&E staining andfluorescent imaging (GFP and DAPI)] as well as SEM.

In vitro Functional Testing Of Re-Endothelialization

To determine the functionality of the re-endothelialized livervasculature, MS1-seeded scaffolds were perfused with freshly collectedheparinized porcine blood for 1 hour. The blood perfusate was preparedby diluting whole blood at a 3:1 ratio in Kreb's bicarbonate buffer.Unseeded scaffolds were used as controls. To determine platelet adhesionon the scaffolds, paraffin sections were stained with an anti-integrinaIIb antibody (Santa Cruz Biotechnology Inc., Santa Cruz, Calif.) andAlexa 647-conjugated, rabbit anti-goat antibody (Invitrogen LifeTechnologies), followed by visualization by confocal microscopy (LSM5,Zeiss). The levels of platelets adhered onto vessel walls werequantified using the confocal images from representative regions of thevasculatures (4 random areas, n=3 livers per group) from within theliver parenchyma by measuring the fluorescence intensity using Image Jsoftware. The fluorescence intensities were averaged, and thenrepresented as mean±SD.

In vivo Implantation Study

Yorkshire pigs (female, 60e80 kg) were used for the in vivo animalstudy. All surgical procedures were performed in accordance with theInstitutional Animal Care and Use Committee (IACUC) approved protocols.Recipient animals were partitioned into two different groups: i)decellularized liver scaffold without further treatments, n=3 animalsand ii) re-endothelialized liver, n=3 animals. The implantation protocolwas similar to that developed in our previous study [27]. Briefly,animals were placed in a supine position. The surgical site wasdisinfected, and an intravenous catheter was placed to allow for fluidreplacement and drug delivery during surgery. Cardiopulmonary functionwas monitored throughout the procedure. Under general anesthesia, amidline abdominal incision was made through the skin and muscle. Theabdominal aorta and inferior vena cava just above the iliac bifurcationwere identified and dissected from the surrounding tissue. After placinga large Satinsky clamp on the renal artery and renal vein, the portalvein and IH-IVC of the liver scaffold were anastomosed to the aorta andinferior vena cava in an end-to-side fashion using double-armed 6/0prolene sutures. After reperfusion, the blood flow through the arteryand vein of the implanted scaffold was monitored with Color Dopplerultrasonography (Acuson Sequola 512, Simens).

Characterization of the Harvested Liver

At 1 day after implantation, vascular patency in the implanted liverscaffold from the animals was examined by ultrasound imaging andradiographic fluoroscopy using Siemens SIREMOBIL Compact L C-arm(Siemens, Munich, Bavaria, Germany). Conray contrast agent (iothalamatemeglumine) (Mallinckrodt Inc., St Louis, Mo.) for the fluoroscopicangiogram was injected through the renal artery. After euthanization,the retrieved liver scaffold was examined by histological (H&E) andimmunohistochemical (platelet immunostaining) analysis. Forquantification of platelet adhesion, aIIb+platelet fluorescenceintensity was obtained from randomly selected images (four per liverfrom the portal vein branch and central vein). Fluorescence intensitywas calculated from the values (total fluorescence intensity/area)] ofeach fluorescent image using Image J software. The data was representedas mean±SD.

Statistics

Student t-test analysis was used for statistical analyses. Differenceswere considered significant at P<0.05.

Preparation and Characterization of Acellular Porcine Liver Scaffolds

Perfusion of the decellularization reagent through the portal vein (PV)and hepatic artery (HA) resulted in a gradual color change to whiteafter 1 day, indicating progressive removal of the cellular componentswithin the liver scaffold (FIG. 33A). To evaluate the efficacy ofdecellularization, histological analysis and DNA quantifications wereperformed (FIG. 33B). Histological analysis showed that thedecellularized scaffold contains no cellular materials at the PV branchand central vein as well as within the liver parenchyma (H&E).Quantification of residual DNA within the liver scaffold indicated asignificant reduction as compared to native liver (P<0.05).

To determine the preservation of intact vasculatures within theacellular liver scaffolds, CT angiographic analysis and vascular castingin combination with scanning electronic microscopy (SEM) were performed(FIG. 33C). CT images of the native and decellularized liver scaffoldsdemonstrate intact vasculatures throughout the lobes of the scaffolds.Additionally, liver vascular casting showed preservation of a functionalvascular network within the scaffolds that is capable of allowingefficient perfusion through the entire construct. Importantly, SEManalysis of the liver vascular casts showed intact lobular structures(arrows) within the native and decellularized liver casts, indicatingmaintenance of an intact capillary bed within parenchymal lobules.

Re-Endothelialization of the Liver Scaffold and Structural andFunctional Characterization

Homogenous and functional re-endothelialized vasculatures within thedecellularized porcine liver scaffolds are critical for establishment ofvascular patency. To enhance the attachment of endothelial cells andprevent detachment under physiologic flow conditions, CD31 antibodieswere conjugated onto the vascular surfaces, as confirmed by imaging of afluorescently labeled secondary antibody. The images clearly showedincreased fluorescence intensity in antibody-conjugated vasculatures ascompared with the unconjugated constructs. This indicated efficientantibody conjugation onto vasculatures of the decellularized scaffolds.The statistical difference was significant in vessels of varyingdiameters (Student t-test, P<0.05) (FIG. 33D). For re-endothelializationof acellular liver scaffolds, murine endothelial cells (MS1) were seededonto the scaffold using a combination of static and perfusiontechniques. Results from fluorescence imaging and SEM analysisdemonstrated that the MS1 cells expressing GFP are intact, well-aligned,and uniformly distributed over the lumen of large vessels (FIG. 34A).Efficient re-endothelialization was also observed on intrahepaticvessels such as PV branches and central veins within portal triads (FIG.34B). Based on the GFP fluorescence imaging, 80-90% of theextracapillary vasculature, including the portal vein (PV), inferiorVena Cava (IVC), PV branches, and central vein (CV) branches, arecovered by endothelial cells.

To determine whether the seeded liver scaffolds are functional,heparinized pig whole blood was perfused through the PV. It was expectedthat the functional re-endothelialization of vasculatures would reduceplatelet adhesion in re-endothelialized liver scaffolds during bloodperfusion. After blood perfusion in vitro, the effect ofre-endothelialization was determined by immunostaining with integrinaIIb antibodies on unseeded and seeded liver matrices. A minimal numberof integrin aIIb+platelets were found on the portal vein and centralvein of the re-endothelialized liver scaffolds [EC(+)], whereas highlevels of platelet accumulation were observed in the EC(−) livers (FIG.34C). Quantitatively, the portal vein and central vein of EC+liversshowed significantly reduced platelet adhesion. This reduction was 3-4fold compared to EC⁻ livers (FIG. 34C, Student t-test, P<0.05).Quantification of platelet adhesion by measurement of fluorescenceintensity showed a significant decrease in platelet adhesion in theseeded scaffolds (FIG. 34C). These results indicate that endothelialcell seeding of the acellular porcine liver scaffold results information of a uniform and functional endothelial layer inside thevascular remnant lumens.

Implantation of Engineered Porcine Liver Construct

To test the feasibility of implanting a clinical-scale bioengineeredliver, liver constructs (FIG. 35A) were heterotopically implanted intopigs by vascular anastomosis following a left nephrectomy (FIG. 35B).The blood flow and vascular patency were determined by ultrasound andangiography. Following implantation, all liver constructs showed goodblood flow within the liver constructs, as confirmed by ultrasoundimages of blood inflow through the renal artery/portal vein (RA/PV)anastomosis (FIG. 35C), outflow through the intrahepatic-inferior venacava/renal vein (IH-IVC/RV) anastomosis (FIG. 35D), and flow through thebranches of the PV within the left lateral (LL) (FIG. 35E) and rightlateral (RL) lobes (FIG. 35F). Results from the fluoroscopic angiographystudies confirmed vascular patency through the PV, left lateral (LL)lobe, and IH-IVC heparinized as indicated by the arrows (FIGS. 35G-I).

At one day post-implantation, results from vascular imaging demonstratedthat the re-endothelialized liver constructs continued to maintain goodblood flow and vascular patency as compared to the unseeded scaffolds(FIGS. 36A-B). Ultrasound imaging (FIG. 36A) of the unseeded scaffolds(EC⁻) showed the absence of blood flow into the liver implant; while theimaging of constructs from the EC⁺ group showed a significant blood flowinto the portal system along with good outflow from the RV.Additionally, a significant blood flow within the liver parenchyma (LLand RL lobe) was noted in the EC⁺ constructs as compared to theacellular scaffolds.

As expected, significant differences in terms of vascular patency wereobserved between the unseeded and re-endothelialized groups (FIG. 36B).The unseeded liver implants showed a complete absence of vascularpatency. During direct injection of contrast media into the portal vein(PV) for fluoroscopic angiography, a very strong resistance was noted,which was likely due to the complete clotting of the entire portalsystem. While flow into some main PV branches was detectable, the entireparenchyma was obstructed by blood clots. In contrast, the constructsfrom the EC⁺ groups demonstrated maintenance of the native geometricalvessel hierarchy as visualized by the injected contrast agent (arrows),indicating that these vascular networks maintained patency. The EC⁻group displayed vascular patency only inside the main portal branches,with irregular and scattered diffusion of the contrast medium within theconstruct. However, constructs from the EC⁺ group showed a homogeneousdistribution of the contrast agent from the hilar zone to peripheralregions of each lobe. The maintenance of vascular patency wasaccompanied by the presence of endothelial cells within the vasculaturesof the implant (FIG. 36C), which resulted in the reduction of plateletadhesion on the vessel walls. Quantification of platelet adhesion showeda significant difference between the EC⁻ and EC⁺ groups (Student t-test,P<0.05).

Production of bioengineered livers for transplantation into patientsundergoing liver failure would have an enormous impact on the healthcarefield, as this technology could alleviate the current problem of donororgan shortage [26]. Successful generation of bioengineered tissues withsimpler structures have been achieved to date, including blood vessels[28], bladder [29] and upper airways segments [30]. However, engineeringof complex, solid organs such as kidney and liver pose severaladditional challenges that must be met before they can be employedclinically. One major obstacle in creating an engineered functionalsolid organ is reestablishment of the organ's vasculature to a degreethat is capable of 1) withstanding physiological blood flow conditions,2) preventing thrombosis, and 3) able to deliver blood to the entireorgan's volume. In the current study, this challenge was overcome withthe results showing that re-endothelialized liver scaffolds were able towithstand physiologic blood pressure and maintain blood flow within thebioengineered livers for 24 hours.

Prevention of thrombosis requires coverage of the vascular lumen byendothelial cells in order to prevent platelet adherence to vessel wall.In the current study, successful re-endothelialization of thevasculatures of decellularized liver matrices was demonstrated (FIG.32). An antibody conjugation technique was employed in order to enhanceendothelial cell adhesion during in vitro maturation of the construct,and following implantation in vivo. In our previous study usingacellular porcine kidney scaffolds [31], we confirmed effectiveness ofCD31 antibody conjugation by demonstrating that antibodyconjugation-mediated endothelial cell attachment on the vasculature ofimplanted scaffolds improved vascular patency throughout the renalmatrix, when compared to the re-endothelialized scaffolds withoutantibody conjugation. In a similar way, for this liver study, theantibody conjugation method was used for better maintenance ofre-endothelialization of the engineered liver construct during in vivoimplantation. As expected, efficient re-endothelialization via the CD31antibody conjugation of bioengineered livers prevented thrombosis undernormal physiological vascular conditions. While reendothelializationusing anti-CD31 antibody conjugation reinforces endothelial cellattachment to the vascular surfaces, it may also be possible toco-conjugate anti-thrombotic agents, such as heparin, to furtherminimize the potential for thrombosis and allow additional time formaturation of the vasculature in vivo.

The porcine transplant model was chosen for this study in order tovalidate the function of the reestablished vasculature under clinicallyrelevant conditions. Furthermore, a large animal model for bioengineeredorgan transplantation represents a significant scale-up of thistechnology from published studies using rodents [19]. In terms of size,our engineered porcine liver constructs measure approximately 15-20 cm(FIG. 33A) in diameter, which is comparable to that of human liver[greatest transverse (20-22.5 cm), vertical (15-17.5), and greatestantero-posterior diameter (10-12.5 cm) measurement] [32]. The combinedhepatic blood flow rate through the portal vein and hepatic artery isapproximately 700 ml/min [33]. Maintaining flow through the transplantedconstruct for 24 hours indicates the potential for handling up to 1000 Lof blood over this time frame. This is the first report thatdemonstrates implantation of a vascularized liver construct ofclinically relevant scale. This finding leads to the expectation thatthe technology will be useful in humans.

Vessel casting of decellularized livers demonstrated that the sinusoidalbasement membrane within the lobular structures is capable of retainingthe casting resin. This finding was further supported by histologicalanalysis that confirmed re-endothelialization extending to the capillarybed of the construct.

These data indicate that complete re-endothelialization of a large-scaleliver construct is possible and has been achieved. Fluoroscopicangiography showed that the level of re-endothelialization achieved inthe current study was sufficient to withstand physiological blood flowconditions for up to 24 hours. This finding is consistent with theDoppler ultrasound data, which measured blood flow within thetransplanted construct throughout the 24-hour period. These cumulativefindings indicate that re-endothelialized liver constructs have thepotential to support a hepatic parenchyma for extended periods of time,in an in vivo model of clinically relevant scale. This findingestablishes that the technology disclosed herein allows for thephysiological perfusion of bioengineered large organs such as liver,kidney and pancreas (see Example 12) in order to ensure viability andfunction of the parenchymal cells. In the current study, seeded liverconstructs were allowed to partially mature in a perfusion bioreactorfor three days. However, further maturation and remodeling of theconstruct would be expected following transplantation in vivo.

Previous studies by our group indicate that decellularized livermatrices are a suitable scaffold to support cell phenotypes found in theliver parenchyma, including mature hepatocytes [19, 34, 35]. However,for these parenchymal cells to remain viable within the construct, anunobstructed blood flow to these cells must be maintained. The datapresented here represent a significant step towards this end. Whilecells isolated from cadaveric livers deemed unsuitable for transplantrepresent one potential parenchymal cell source, a construct engineeredusing these cells would still carry with it the requirement forimmunosuppression following transplant. Recent advances in the field ofinduced pluripotent stem cells (iPS) provide a potential autologous cellsource for whole-organ engineering. These cells have also been shown topossess significant capacity for in vitro expansion, meaning that anumber of cells sufficient for construct seeding may be obtained from asmall population of iPS derived hepatocytes [36].

In conclusion, we have demonstrated for the first time the feasibilityto generate natural porcine liver ECM scaffolds with an intact vascularnetwork extending to the capillary bed, which is sufficient to preventthrombosis within the liver construct for a period of 24 hours followingtransplantation. Furthermore, we have accomplished these advances inpigs, which represent a clinically relevant scale. These findingsdemonstrate the feasibility of engineering a whole liver vasculaturethat would allow integration into the circulatory system to support abioengineered liver long-term. We see this as a necessary first steptowards the generation of an engineered liver for the treatment of endstage liver disease.

REFERENCES—FOR EXAMPLE 11 ONLY

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Whole-organ bioengineering: current tales of modern alchemy.        Transl Res 2014; 163: 259-67.    -   [10] Faulk D M, Johnson S A, Zhang L, Badylak S F. Role of the        extracellular matrix in whole organ engineering. J Cell Physiol        2014; 229:984-9.    -   [11] Daley W P, Peters S B, Larsen M. Extracellular matrix        dynamics in development and regenerative medicine. J Cell Sci        2008; 121:255-64.    -   [12] Schenke-Layland K, Rofail F, Heydarkhan S, Gluck J M, Ingle        N P, Angelis E, et al. The use of three-dimensional        nanostructures to instruct cells to produce extracellular matrix        for regenerative medicine strategies. Biomaterials 2009;        30:4665-75.    -   [13] Ott H C, Clippinger B, Conrad C, Schuetz C, Pomerantseva I,        Ikonomou L, et al. Regeneration and orthotopic transplantation        of a bioartificial lung. Nat Med 2010; 16:927-33    -   [14] Ott H C, Matthiesen T S, Goh S K, Black L D, Kren S M,        Netoff T I, et al. Perfusiondecellularized matrix: using        nature's platform to engineer a bioartificial heart. Nat Med        2008; 14:213-21.    -   [15] Peloso A, Katari R, Zambon J P, Orlando G. Sisyphus, the        Giffen's paradox and the holy grail: time for organ        transplantation to transition toward a regenerative        medicine-focused type of research. Expert Rev Clin Immunol 2013;        9:883-5.    -   [16] Petersen T H, Calle E A, Zhao L, Lee E J, Gui L, Raredon M        B, et al. Tissue-engineered lungs for in vivo implantation.        Science 2010; 329:538-41.    -   [17] Uygun B E, Soto-Gutierrez A, Yagi H, Izamis M L, Guzzardi M        A, Shulman C, et al. Organ reengineering through development of        a transplantable recellularized liver graft using decellularized        liver matrix. Nat Med 2010; 16:814-20.    -   [18] Orlando G, Soker S, Stratta R J. Organ bioengineering and        regeneration as the new holy grail for organ transplantation.        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Biochim Biophys        Acta 2014; 1840:2506-19.    -   [25] Yagi H, Soto-Gutierrez A, Kitagawa Y. Whole-organ        re-engineering: a regenerative medicine approach to digestive        organ replacement. Surg Today 2013; 43:587-94.    -   [26] Soto-Gutierrez A, Wertheim J A, Ott H C, Gilbert T W.        Perspectives on whole organ assembly: moving toward        transplantation on demand. J Clin Invest 2012; 122:3817-23.    -   [27] Orlando G, Farney A C, Iskandar S S, Mirmalek-Sani S H,        Sullivan D C, Moran E, et al. Production and implantation of        renal extracellular matrix scaffolds from porcine kidneys as a        platform for renal bioengineering investigations. Ann Surg 2012;        256:363-70.    -   [28] Hoerstrup S P, Cummings Mrcs I, Lachat M, Schoen F J, Jenni        R, Leschka S, et al. Functional growth in tissue-engineered        living, vascular grafts: follow-up at 100 weeks in a large        animal model. Circulation 2006; 114:1159-66.    -   [29] Atala A, Bauer S B, Soker S, Yoo J J, Retik A B.        Tissue-engineered autologous bladders for patients needing        cystoplasty. Lancet 2006; 367:1241-6.    -   [30] Macchiarini P, Jungebluth P, Go T, Asnaghi M A, Rees L E,        Cogan T A, et al. Clinical transplantation of a        tissue-engineered airway. Lancet 2008; 372:2023-30.    -   [31] Ko I K, Abolbashari M, Huling J, Kim C, Mirmalek Sani S H,        Moradi M, et al. Enhanced re-endothelialization of acellular        kidney scaffolds for whole organ engineering via antibody        conjugation of vasculatures. Technology 2014; 2:243-53.    -   [32] Gray H. The liver. 2i. In: Anatomy of the human body. 20th        ed. 1918. p. 1-54.    -   [33] Winterdahl M, Sorensen M, Keiding S, Mortensen F V, Alstrup        A K, Hansen S B, et al. Hepatic blood perfusion estimated by        dynamic contrast-enhanced computed tomography in pigs:        limitations of the slope method. Invest Radiol 2012; 47:588-95.    -   [34] Shupe T, Williams M, Brown A, Willenberg B, Petersen B E.        Method for the decellularization of intact rat liver.        Organogenesis 2010; 6:134-6.    -   [35] Skardal A, Smith L, Bharadwaj S, Atala A, Soker S, Zhang Y.        Tissue specific synthetic ECM hydrogels for 3-D in vitro        maintenance of hepatocyte function. Biomaterials 2012;        33:4565-75.    -   [36] Zhang R, Takebe T, Sekine K, Koike H, Zheng Y, Taniguchi H.        Identification of proliferating human hepatic cells from human        induced pluripotent stem cells. Transplant Proc 2014; 46:1201-4.    -   [37] Crapo P M, Gilbert T W, Badylak S F. An overview of tissue        and whole organ decellularization processes. Biomaterials 2011;        32:3233-43.    -   [38] Arruebo M, Valladares M, Gonzalez-Fernandez A.        Antibody-conjugated nanoparticles for biomedical applications. J        Nanomater 2009; 2009:1-24.    -   [39] Arbiser J L, Moses M A, Fernandez C A, Ghiso N, Cao Y,        Klauber N, et al. Oncogenic H-ras stimulates tumor angiogenesis        by two distinct pathways. Proc Natl Acad Sci U. S. A 1997;        94:861-6.

EXAMPLE 12 Whole Organ Engineering—Re-Endothelialization of PancreasScaffolds

Diabetes mellitus (DM) has reached pandemic levels and represents agrowing burden both on health-care expenditures as well as the qualityand quantity of life for afflicted individuals (194). First-linetreatment is medical (oral agents and insulin) and behavioral (dietaryrestrictions and physical activity). Although exogenous insulin therapyis effective at preventing acute metabolic decompensation and islife-saving for type 1 DM (T1DM), less than 40% of patients achievetherapeutic goals (5). Overall, a large number of diabetic patients areinadequately controlled, complete and steady remission of hyperglycemiasare rare, and treatment may be complicated by hypoglycemia, with mostpatients developing irreversible organ complications during theirlifetimes. This means that standard exogenous insulin-based intensiveglucose control can significantly reduce, but not completely protectagainst, the long-term complications of DM. β9cell replacement througheither islet or pancreas transplantation (PTX) are the only therapiesable to reliably re-establish a stable, long-term euglycemic state. PTXis indicated for patients with T1DM and some selected cases of type 2 DM(T2DM), yet it remains underutilized. In fact, in the US, for every10,000 patients with T1DM, only 3 will actually receive PTX or islettransplant in their lifetime, due to factors such as the lack ofsuitable pancreas donors, the burden of chronic toxicity determined bylife-long immunosuppression, and other issues related to financing andaccess to transplantation (5). When T2DM is considered, only threepatients in one million will ever receive PTX or islet transplant.Therefore, the identification of a new, potentially inexhaustible sourceof β-cells for transplantation is extremely urgent.

Regenerative medicine has shown an immense potential to address thelimited number of transplantable organs and to allowimmunosuppression-free transplantation (9-18). Among the differenttechnologies under development, the seeding of cells on supportingscaffolding material (cell-on-scaffold seeding technology, CSST) (19)appears to offer the quickest route to clinical application. Thistechnology has allowed the production of relatively simple organs thatwere eventually implanted in more than 200 patients, for therapeuticpurposes (reviewed by Orlando) (9). As these body parts werebioengineered from patient's own cells, immunosuppression was neverneeded. CSST is also being considered for bioengineering more complexorgans for transplant purposes. We have developed an innovative methodthat, through the detergent-based de-cellularization of organs (e.g.,porcine and human), allows successful and consistent production ofacellular extracellular matrix (ECM) scaffolds (20-22). These scaffoldsrepresent a biochemically, geometrically and spatially suitable platformfor bioengineering investigations because they are biocompatible, havebasic structural components well-preserved, retain essentialmatrix-bound growth factors and cytokines, and are able to drivedifferentiation of progenitor cells into an organ-specific phenotype.Moreover, the framework of the innate vasculature remains intact and,when scaffolds are implanted in vivo, the framework has the ability tosustain physiological blood pressure. Initial promising investigationsin kidney and endocrine pancreas bioengineering have allowed productionof natural scaffolds through the de-cellularization of porcine and humankidneys, as well as of porcine pancreata. In this latter case, awhole-organ, three-dimensional, acellular scaffold was generated usingthe porcine pancreas. Histology and imaging studies confirmed that theprotocol disclosed herein effectively removes cellular material whilepreserving ECM proteins and the framework of the native vascular tree.Moreover, when these scaffolds are seeded with precursor cells, cellsadhere, proliferate and are both viable and functioning; when pancreaticislets are seeded, they are able to respond to a glucose stress test(20).

It is expected that when a patient will need beta cell replacementtherapy, s/he will be offered a new generation bio-artificial endocrinepancreas (BAEP) produced from his/her own cells that will be seeded inthe three-dimensional framework of ECM scaffolds manufactured frompancreata of human or porcine origin. Therefore, we have applied thedisclosed de-cellularization method to discarded human pancreata inorder to generate acellular pancreatic ECM scaffolds. Disclosed hereinare investigations on the generation of human pancreas acellular ECMscaffolds (hpaECMs), which represent the hardware of the bioengineeredBAEP.

Pancreas and Endothelial Cell Procurement

Twenty-five human pancreata obtained from the service area of our localprocurement organization (Carolina Donor Services—CDS) were processed.Each organ was originally recovered for transplantation, butsubsequently discarded for steatosis, fibrosis, vascular anomalies orinflammation. After the national list was exhausted, pancreata fromdonors with research consent were released for subsequentinvestigations.

Pancreas Preparation

Organs were received in cold sterile preservation solution after en-blocremoval that included the whole organ pancreas, duodenum and spleen, andthen prepared by removing the duodenum and ligating all of the vascularbranches from the arterial arcade connecting the pancreatic head withthe duodenum using 390 silk sutures. Thereafter, the pancreatic duct(PD) was cannulated with a 14 G catheter. The peripancreatic fat wastrimmed, the splenic vessels were ligated at the splenic hilum, and thespleen was removed. The superior mesenteric artery (SMA) and proximalstump of the splenic artery (SA) were cannulated using 16 G plasticconnectors. Pancreata were then washed with 500 ml of saline solutioncontaining 50 ml of betadine and then with 10% Penn/Strep solution.Pancreata were eventually rinsed with 500 ml of sterile PBS and storedat 4° C.

Pancreas Decellularization

After placing the pancreas in an ad hoc container, PD, SMA and SA wereconnected to a double-line peristaltic pump (Masterflex L/S, easy loadpump head and L/S 16G tubing, Cole-Parmer Instrument Co., Vernon Hills,Ill., USA) (FIG. 1). Then, the pancreas was flushed with 4° C. phosphatebuffer solution (PBS) and heparin (1% of 1000 U, 10 U/ml) at 6 ml/minutefor 60 minutes (360 mL total). Afterwards, 1% Triton X-100 and 0.1%ammonium hydroxide solution were perfused at 12.5 ml/minute for 48 hours(72 L total) through both inlets (PD and SMA/SA inlets), at 4° C. Theso-obtained hpaECMs were then rinsed with DNAse (D5025-Type IV,deoxyribonuclease from bovine pancreas, Sigma Aldrich, St. Louis, Mo.)and 0.0025% magnesium chloride). Finally, scaffolds were perfused withsaline at a flow of 6 mL/minute for 5 days (12,900 L total) to removedetergent and stored at 4° C. Scaffolds destined for cell-seedingexperiments (see below) were sterilized by γ-irradiation (12,000 Gy) andstored in PBS at 4° C.

Bioscaffold Characterization—Vessel Patency

To evaluate the patency of the hpaECMs' innate vasculature, angiographywas performed by injecting 5 ml of contrast agent in each of the inlets(PD, MSA and SA, 15 mL total) at a flow rate of 25 mL/minute.

Bioscaffold Characterization—Collagen and DNA Quantification

Collagen and DNA content of fresh and de-cellularized pancreas wasmeasured through indirect quantification of hydroxyproline residuesusing the Quick9Zyme Total Collagen Assay (QuicZyme, Biosciences), andusing a tissue DNA isolation kit (PureLink Genomic DNA MiniKit,Invitrogen), respectively, as described by Orlando et al. (22).

Bioscaffold Characterization—Basic Histology, Immunofluorescence andCellularity

hpaECMs samples were fixed for 24 hours in formalin, washed in diH2O,dehydrated in graded alcohol, embedded in paraffin, 5 mm-sectioned, andeventually stained with H&E, Masson's Trichrome (MT), elastic Van Gieson(EVG), Alcian Blue (AB) and Picrosirius Red (PR). Slides were comparedwith counterparts obtained from discarded pancreata that did not undergode-cellularization.

For immunofluorescence analysis, slides were incubated overnight withprimary antibodies against collagen type I (dilution 1:25, SouthernBiotech), collagen type IV (dilution 1:100, Southern Biotech),fibronectin (dilution 1:100, Santa Cruz), laminin (dilution 1:200,Sigma), HLA Class I (dilution 1:100, Abcam) and HLA DP+DR+DQ (dilution1:25, Abcam). Nuclei were counterstained with DAPI. Slides werevisualized using fluorescence microscopy (Carl Zeiss) and compared tocounterparts obtained from discarded pancreas that did not undergode-cellularization.

To quantify the remaining cells after de-cellularization, a portion offresh and acellular tissue was frozen in liquid nitrogen and 10 slidesof each tissue were analyzed with an optical microscope. The sampleswere covered with Vectashield (Vector Laboratories, Inc., Burlingame,Calif.) mounting medium for fluorescence with40-6-diamidino-2-phenylindole (DAPI) (Vector Laboratories, Inc), and thetotal number of nuclei per field was counted in random pictures usingfluorescence microscopy (Carl Zeiss).

Bioscaffold Characterization—Scanning Electron Microscopy (SEM)

To evaluate scaffold ultrastructure, samples of hpaECMs and normalpancreas were processed and analyzed as described by Orlando et al.(22). Images were recorded with a Jeo17401 FEG scanning electronmicroscope.

Bioscaffold Characterization—Chicken Chorioallantoic Membrane (CAM)Angiogenic Assay

To assess the angiogenic properties of the scaffolds, a CAM assay wasused, as previously described (22-24). To assess angiogenesis, thenumber of blood vessels converging towards de-cellularized matrices wascounted manually. Only blood vessels less than 10 mm in diameter werecounted by blinded assessors (n=4), with the mean of the counts beingconsidered.

Bioscaffold Characterization—Measurement of Pancreatic Tissue Stiffness

To assess hpaECMs stiffness, dog bone-shaped samples, 4 mm wide at thenarrowest point with a length of 18 mm, were punched from wet pancreasbefore and after the de-cellularization process. The mechanicalproperties were measured using tensile extension by a uniaxial load testmachine (Instron5544, Instron Corporation, Issaquah, Wash.) with anextension rate of 10.0 mm/minute. Stress-strain curves for scaffoldswere recorded and Young's modulus, tensile strength, and strain at breakwere obtained. Six samples were measured for each scaffold allowing themean and standard deviation to be calculated.

Bioscaffold Characterization—Growth Factor (GF) Analysis

We expected growth factors, cytokines and chemokines to be onlypartially depleted from our matrices. To confirm this expectation, 5hpaECMs were produced from 5 different organs (study group). Two sampleswere procured from each scaffold (10 pieces total) with a 12 mm biopsypunch; the control group consisted of 10 samples obtained in the samefashion from 5 discarded pancreata that did not undergode-cellularization. All samples were stored in sterile PBS with 2%pen/strep and then shipped to the manufacturer (Raybiotech,http://www.raybiotech.com), where they were processed using theQuantibody® Human Growth Factor Array Q1. The results of the experimentsassessing depletion of growth factors from the matrices are presented inTable 3.

TABLE 3 Angiogensis and vasculogenesis Embryogenesis, growth andregeneration N

P

N

P

Growth factor

Growth factor

EG-VEGF 28.33 25.

8

P

7

3 83.21 V

3.34 289.9

4

.3

HB-EGF

.28

.78 37.82

34.89 T

.22

51.

4

1 13.71

2

11.

AR

8.

S

113.

59.

82162.

.13 3

SC

.15

48.

37.17

.2

31

.23

69.

2 3.81

D

Viscero afferent neuronal development Insulin level and glucosehomeostasis control N

h

M P

N

M P

Growth factor

d

Growth factor

S

13.

GM

8

NGF 4.18 0.5

.38

3

7

4.

NT-3

2 2.8 18.31

8.

7

8

NT-4

2

H

3.8

5

.17 3.

1.87

indicates data missing or illegible when filed

GF analysis. Table 3 illustrates the growth factors (GFs) retainedwithin hpaECMs after decellularization. GFs were divided in four groupsby their function: angiogenesis and vasculogenesis (Group 1), growth andregeneration (Group 2), neuronal development (Group 3) and insulin leveland glucose homeostasis Group 4). In Group 1, EG-VEGF (EndocrineGland—derived Vascular Endothelial Growth Factor) and VEGF9r3 (VascularEndothelial Growth Factor receptor 3) were significantly present (87.35%and 40.34% of the initial content, respectively); VEGF-r2 (vascularendothelial growth factor receptor 2), AR (amphiregulin),FGF94(fibroblast growth factor 4), bFGF (fibroblast growth factorbinding protein) were moderately present (13.71%, 12.63%, 14.09% and10.95%, respectively) whereas VEGF (vascular endothelial growth factor),FGF97 (fibroblast growth factor 7) and VEGF D (vascular endothelialgrowth factor D) were poorly present (9.2%, 3.81% and 0,97%,respectively) (Table 3). In Group 2, PDGF-AA (platelet-derived growthfactor AA), HB-EGF (heparin binding epidermal growth factor), TGFα(transforming growth factor α) and TGFβ1 (transforming growth factor β1)were well-preserved at 81.21%, 57.35%, 48.88% and 49.16% of the initialcontent. MCFr (mitochondrial carrier family receptor), SCF (stem cellfactor), and TGFβ3 (transforming growth factor β3) were moderatelypresent: 26.05%, 17.17%, and 14.72%, respectively. In Group 3, BDNF(brain derived neurotrophic factor), b-NGF (nerve growth factor bindingprotein), GDNF (glial cell line derived neurotrophic factor) and NT-3were present at 13.97%, 14.18%, 16.09% and 10.11% of the initialcontent, respectively. NT-4 (neurotrophin 4) was almost completelyabsent post-de-cellularization (5.07%). In Group 4, GH (growth hormone),BMP7 (bone morphogenetic protein 7), BMP5 (bone morphogenetic protein5), BMP4 (bone morphogenetic protein 4), IGF-1 (insulin-like growthfactor 1) and NGFr (nerve growth factor receptor) remained at 37.76%,24.7%, 32.6%, 9.23%, 2.6%, and 7.67% of the initial content,respectively.

Assessment of the Ability of hpaECMs to Sustain Cell Attachment andGrowth

a) Human islets: Pancreases were processed using standard Liberase HIcollagenase digestion with mechanical dissociation within a Ricordichamber. Islets were purified with an optiprep discontinuous gradientmethod, identified and counted using dithizone staining. paECMs sampleswere prepared as shown in FIG. 37, sterilized via γ-irradiation andplaced in RPMI with 10% FBS overnight at 4° C., prior to seeding. Theday after, 200 islets per scaffold were statically seeded on matriceswith fresh media and cultured under standard conditions in RPMI mediawith 10% FBS for 4 days prior to perifusion testing (20). As control,additional islets were cultured on standard Petri dishes under the sameculture conditions. Briefly, each group of 200 islets was placed intoperifusion chambers, either seeded on scaffolds or free. The chamberswere then perfused with Krebs-Ringer-bicarbonate (KRB) solutioncontaining 0.2% bovine serum albumin (BSA) at a flow rate of 0.3mL/minute. During the procedure, the KRB was continuously gassed with95% air/5% CO₂. The islets were pre-perfused for 1 hour in a low-glucose(3.3 mM) KRB solution prior to sampling. After one hour, effluentsamples were collected from the low-glucose KRB solution every 5 minutesfor 20 minutes. Then a high-glucose solution (16.7 mM) was perfusedthrough the chambers for 30 minutes with effluent sample collectionevery 2 minutes. Following the 30 minutes of high glucose, the chamberswere perfused again with low-glucose KRB for 30 minutes with samplecollection every 5 minutes. The effluent samples were stored at −20° C.until analyzed for insulin content via radioimmunoassay. Insulin valuesare normalized to basal averages, n=4, error bars represent SEM.

b) Human endothelial cells: Human primary pancreatic endothelial cells(hPPECs) were isolated from discarded pancreata as described by Navoneet al. (25). Cells were then expanded on matrigel-coated flasks and grewwith endothelial proliferation medium (EndoPM). At the time of seeding,20×10⁶ cells were injected with a syringe pump in a whole acellularpancreas scaffold through the SMA and SA, at a flow of 0.2 ml/minute.Cells were allowed to attach for 2 hours, after which perfusion culturewas started. Seeded matrices were cultured in a custom-built bioreactor(FIG. 43B). Media were infused through Luer lock access ports andallowed to equilibrate with 5% CO₂ and 95% room air by insertion of 0.22μm filters and magnetic stirring. The system (and the related technicalsupport) was designed and developed by SKE Advanced Therapies. After 6days, matrices were fixed in formalin, sampled into 5-μm sections, andstained for H&E, CD31 (clone JC70A, 1:100, DAKO), Ki67 (clone Mib1,1:600, DAKO) using standard protocols. Antigen retrieval was performedwith Tris EDTA pH 9 for both antibodies. The percentage of Ki67+ cellswas obtained as the ratio of Ki67+ and hematoxylin+ cells.Immunohistochemistry and H&E-stained images were recorded using AperioScan System (Leica Microsystems).

Human Pancreas De-Cellularization

Triton-based perfusion allowed complete clearance of the cellularcompartment of discarded pancreata (FIGS. 37A-B). Fluoroscopy depictedan intact vascular network that retained the hierarchical branchingstructures (Supplementary FIGS. 37C-G). In fact, contrast media flowedprogressively from larger vessels to smaller capillaries to eventuallydrain out through the venous outlets, without extravasating within thehpaECMs.

Collagen and DNA Quantification

To investigate the ability of the de-cellularization protocol tomaintain ECM components of the native pancreas, fresh and acellularsamples were processed for collagen quantification. Quantitative assayshowed statistically increased collagen in acellular matrices comparedto fresh tissue (from 22.9±10.0 to 75.4±16.8 μg/mg, P<0.001). DNAanalysis was performed as indirect proof of successful decellularizationand demonstrated approximately 99% clearance of DNA in comparison withthe native organ (from 1.35±0.43 to 0.04±0.03 μg/mg, P<0.001) (FIGS.1C-G).

Basic Histology, Immunofluorescence and Cellularity

Fresh and acellular samples were processed for Masson trichrome,Picrosirius red, elastic Van Gieson and Alcian blue (FIG. 38). Allstainings confirmed complete absence of nuclear material and cells,consistent with the H&E and DAPI findings. Masson trichrome andPicrosirius red revealed preservation of collagen fibers, both in thenative tissue and acellular matrices. Alcian blue staining indicatedsome preservation of sulfated glycosaminoglycan proteins in thescaffolds. Elastic Van Gieson staining showed that elastin was retainedafter de-cellularization. Elastin is an essential component of the basalmembrane of vascular structures and ducts in the pancreas, ashighlighted by EVG staining in fresh samples.

DAPI and H&E staining, as well as immunofluorescence for both MHC classI and class II antigens confirmed the complete clearance of the cellularcompartment of the hpaECMs (FIG. 39). As expected, the scaffoldingarchitecture of the pancreas was well preserved. In fact, H&E stainingshowed the pink eosinophilic staining typical of collagen in the absenceof any basophilic staining indicative of cellular nuclear material.Specific nuclei staining with DAPI confirmed the absence of cell nucleiafter de-cellularization. This finding was also supported by the countof the total number of nuclei per field in random pictures (from 410±120to 0 nuclei, P<0.001). As an indirect sign of successfulde-cellularization, neither MHC class I- nor class II-positive cellswere seen in the pancreatic matrices.

Immunofluorescence for specific ECM molecules that have a significantrole in the pancreatic parenchyma was performed for a fullercharacterization of our scaffolds. Immunostaining detected the presenceof laminin, collagen I, collagen IV and fibronectin in the nativepancreas (FIG. 39). Laminin detection in the acellular matrix showedstrong positivity similar to fresh tissue, especially around vessels andstructures resembling the native sites of the islets. In contrast,preservation of collagen I was less evident with respect to the freshpancreatic ECM, while immunofluorescence for collagen IV and fibronectinshowed specific distribution of both components at the same level ofnative tissue.

Scanning Electron Microscopy (SEM) Analysis

SEM revealed retained nanofibrous structures in hpaECMs and confirmedthe absence of intact cells, indicating that the de-cellularizationprocess produced an acellular matrix scaffold that retained the micro-and ultra-structural details of pancreas architecture (FIG. 40).

CAM Assay

To test the ability of pancreatic acellular matrices to induceneo-angiogenesis, fragments of de-cellularized pancreas were placed onthe chicken egg CAM. Macroscopic observations of CAM treated withacellular matrices showed that the scaffolds integrated well with thedeveloping environment of the chicken egg and representative images ofscaffold placed in ovo at 0 and 7 days of incubation demonstratedattraction of blood vessels that, in a spoke-wheel pattern, seem topenetrate the tissue (FIG. 41). The effect of pancreatic acellularmatrices on directed blood vessel growth was quantified by manuallycounting the total number of blood vessels converging towards matrices.Seven days after implantation, the number of vessels growing towards theacellular scaffold was significantly increased in comparison to the samesample at day 0 (P<0.05) and to the filter membrane loaded with PBS thatwas used as a negative control (P<0.05). Interestingly, acellularmatrices pro-angiogenic effect after 7 days from implantation into theCAM, was comparable with VEGF-loaded membrane used as positive control(P>0.05).

Acellular Scaffold Stiffness Properties

It has been proposed that the ECM has the ability to maintain theoriginal mechanical properties of native organs. By assessing themechanical properties of our scaffolds, we were able to partiallyelucidate the influence of the de-cellularization process on ECM interms of destruction, degradation and/or decomposition. The propertiesof native pancreas were compared with that of hpaECMs. The resultsshowed that the Young's moduluses and tensile strain at break had slightdecreases, but no significant difference after the removal of cellularcomponents, changing from 0.165±0.039 MPa and 96.71±39.67% to0.139±0.052 MPa and 76.22±22.23%, respectively (FIG. 44). The tensilestress at break of decellularized pancreata (0.070±0.017MPa) wascomparable to the native pancreas (0.068±0.006MPa) (P<0.05). Theseresults indicated that the ECM plays a prominent role in determining themechanical properties of the pancreas, and that our decellularizationmethod effectively maintains the integrity of the tissue's ECM withoutdestruction or degradation.

hpaECMs Retain GFs

Data obtained from the quantitative array platform showed that severalGFs were still present in the hpaECMs after de-cellularization. Some ofthese molecules are essential in angiogenesis and vasculogenesis, celldevelopment, neuronal growth and regeneration, as well as insulin leveland glucose homeostasis (Table 3).

Cell Seeding of Acellular Pancreatic Scaffold

a) Human Islets: Islets stained positive for DTZ on both scaffolding(FIG. 42D) and culture dishes (FIG. 42C). Both islets seeded onscaffolds and islets on culture dishes were functional following 4 daysof culture with similar stimulation indices (as defined as the peakinsulin secretion rate divided by the basal insulin secretion rate) of2.81±0.48 and 3.44±1.821, respectively (p=0.59, average±standarddeviation, n=3).

b) Human endothelial cells: At H&E, cells were ubiquitous within thematrices, yet tended to localize at the level of small and large vessels(FIG. 43). Cells distributed in a line on the edges of the vessel wallsresembling primitive structures of vessel formation. All cells confirmedtheir endothelial nature as evidenced by CD31 staining. More than 50% ofcells were proliferating, as determined by Ki67 staining, demonstratingthat a mid-term culture in the acellular scaffold allows cell survivaland proliferation.

The experiments disclosed in this Example and elsewhere throughout theapplication represent a progression that supports the expectation thatECM scaffolds generated from native pancreas tissue can serve as aplatform for insulin-producing bioengineered tissue (4, 20, 26). Theimportance of transitioning to human organs is obvious in the context ofclinical relevance. The results disclosed herein indicate that wholediscarded human pancreata can be consistently and successfullydecellularized through detergent-based perfusion methodology. Indeed,cell clearance is achieved despite preservation of the ECM'sarchitecture, composition and mechanical properties. In addition, theframework of the vascular network remains intact at all hierarchicallevels. As a corollary to the complete absence of HLA class I and IIantigens, it may be inferred that the so-obtained scaffolds may not besignificantly immunogenic. Interestingly, for the first time it isdisclosed that numerous GFs are retained in significant amounts withinthe three-dimensional structure of hpaECMs. Some of these GFs are keyplayers in essential pathways such as angiogenesis, cell proliferationand glucose metabolism, and their presence may justify the scaffold'sability to induce angiogenesis in the CAM assay.

The data obtained with discarded human pancreas are consistent with ourprevious experience with discarded human kidneys (22). However, incontrast to that experience, we used Triton rather than the moreaggressive SDS for two reasons: first, because the density of thepancreas texture is inferior to the kidney, which is basicallyrefractory to nonionic detergents such as Triton; second, we believethat pancreatic enzymes released during cell breakdown occurring duringthe de-cellularization process can, in turn, contribute to furtherclearance of the cellular compartment. Moreover, these experiments withhuman pancreas are consistent with experiments performed using porcinepancreas (20). In those experiments, pig pancreata were subjected to a24-hour cycle of de-cellularization with a Triton-based solution.Nevertheless, in the case of discarded pancreata, a 48-hourde-cellularization cycle was used—twice the length—that was infusedthrough three different inlets (PD, SMA and SA), rather than just two,namely the PD and the superior mesenteric vein. The adjustment followedinitial unsuccessful attempts while using only the original two inlets,24-hour approach, in which a frank impairment of the de-cellularizationprocess was observed, possibly attributable to the significant tissuedamage that had justified discard. Overall, it was expected that the3-inlet approach—namely, PD, SMA and SA—would lead to successfulde-cellularization, as it would allow ad hoc delivery of the detergentthroughout the whole pancreas; and that the higher volume would possiblycircumvent hurdles deriving from organ damage such as steatosis andfibrosis.

In order to assess the cytocompatibility of hpaECMs, islets and a uniqueendothelial cell line isolated from the human pancreas, namely hPPECs,were used. In both cases, findings show that the matrices are both andcell-friendly, as they allow attachment, function and maintain theinitial phenotype. In the attempt to assess cytocompatibility, while thechoice of islets is obvious, hPPECs are justified because it is expectedthat the first step to follow the production of hpaECMs will be thereconstitution of the endothelium starting from a patient's ownendothelial cells. Consistent with the disclosures herein relating torenal ECM scaffolds produced from discarded kidneys, it is important toemphasize that the strategy of using discarded organs introduces a newtype of biomaterial, represented by natural scaffolds obtained from themanipulation of organs with pre-existing damage.

REFERENCES—FOR EXAMPLE 11 ONLY

-   -   1) http://www.cdc.gov/diabetes/pubs/estimates11.htm    -   2) Boyle J P, Thompson T J, Gregg E W, Barker L E, Williamson        D F. Projection of the year 2050 burden of diabetes in the US        adult population: dynamic modeling of incidence, mortality, and        prediabetes prevalence. Popul Health Metr 2010; 8:29    -   3) American Diabetes Association. Economic costs of diabetes in        the U.S. in 2012. Diabetes Care 2013; 36:1033946    -   4) Orlando G, Gianello P, Salvatori M, Stratta R J, Soker S,        Ricordi C, Domínguez-Bendala J. Cell replacement strategies        aiming at reconstitution of the beta cell compartment in type 1        diabetes. Diabetes 2014; 63(5):1433944    -   5) Orlando G, Stratta R J, Light J. Pancreas transplantation for        type 2 diabetes mellitus. Curr Op Org Transpl 2011; 16 :1109115    -   6) American Diabetes Association. Diagnosis and classification        of diabetes mellitus. Diabetes Care 2009; 32 (Suppl 1):S62-S67    -   7) Esmatjes E, Fernandez C, Rueda S, Nicolau J, Chiganer G,        Ricart M J, Junca E, Fernandez-Cruz L. The utility of the        C9peptide in the phenotyping of patient candidates for pancreas        transplantation. Clin Transplant 2007; 21:358-362    -   8) Wilkin T J. The accelerator hypothesis: a review of the        evidence for insulin resistance as the basis for type I as well        as type II diabetes. Int J Obes (Lond) 2009; 33:716-726    -   9) Orlando G, Soker S, Stratta R J. Organ bioengineering and        regeneration as the new Holy Grail of organ transplantation Ann        Surg. 2013 August; 258(2):221932.    -   10) Orlando G, Wood K J, De Coppi P, Baptista P M, Binder K W,        Bitar K N, Breuer C, Burnett L, Christ G, Farney A, Figliuzzi M,        Holmes J H IV, Koch K, Macchiarini P, Mirmalek9Sani S H, Opara        E, Remuzzi A, Rogers J, Saul J M, Seliktar D S,        Shapira9Schweitzer K, Smith T, Solomon D, Van Dyke M, Yoo J J,        Zhang Y, Atala A, Stratta R J, Soker S. Regenerative medicine as        applied to general surgery. Ann Surg 2012; 255:8679880.    -   11) Orlando G. Immunosuppression9free transplantation        reconsidered from a regenerative medicine perspective. Exp Rev        Clin Immun 2012:8:1799187.    -   12) Orlando G, Wood K J, Soker S, Stratta R J. How regenerative        medicine may contribute to the achievement of an        immunosuppression9free state. Transplantation 2011; 92(8):e3698.    -   13) Orlando G. Transplantation as a subfield of regenerative        medicine. An interview by Lauren Constable. Expert Rev Clin        Immunol 2011; 7:1379141.    -   14) Orlando G, Wood K J, Stratta R J, Yoo J, Atala A, Soker S.        Regenerative medicine and organ transplantation: Past, present        and future. Transplantation 2011; 91:131097.    -   15) Orlando G, Baptista P, Birchall M, Di Coppi P, Farney A,        Opara E, Rogers J, Seliktar D, Shapira-Schweitzer K, Stratta R        J, Atala A, Wood K J, Soker S. Regenerative medicine as applied        to solid organ transplantation: current status and future        development. Transpl Int 2011; 24:2239232.    -   16) Badylak S F, Weiss D J, Caplan A, Macchiarini P. Engineered        whole organs and complex tissues. Lancet 2012; 379(9819):943952.    -   17) Badylak S F, Taylor D, Uygun K. Whole9organ tissue        engineering: decellularization and recellularization of        three-dimensional matrix scaffolds. Ann Rev Biomed Eng 2011;        13:27953.    -   18) Soto-Gutierrez A, Wertheim J A, Ott H C, Gilbert T W.        Perspectives on whole9organ assembly: moving toward        transplantation on demand. J Clin Invest 2012; 122(11):3817923.    -   19) Salvatori M, Peloso A, Katari R, Zambon J P, Soker S,        Stratta R J, Orlando G. Semi-xenotransplantation: combining        transplantation with organ bioengineering and regeneration        technologies to manufacture complex modular organs.        Xenotransplantation, Jul. 8. 2014. doi: 10.111/xen.12122.    -   20) Mirmalek9Sani A, Orlando G, McQuilling J, Pareta R, Mack D,        Salvatori M, Farney A C, Stratta R J, Atala A, Opara E C,        Soker S. Porcine pancreas extracellular matrix as a platform for        endocrine pancreas bioengineering. Biomaterials 2013;        34(22):5488995.    -   21) Orlando G, Farney A, Sullivan D C, AbouShwareb T, Iskandar        S, Wood K J, Atala A, Stratta R J, Yoo J J, Soker S. Production        and implantation of renal extracellular matrix scaffolds from        porcine kidneys as a platform for renal bioengineering        investigations. Ann Surg 2012; 256(2):363970.    -   22) Orlando G, Booth C L, Wang Z, Totonelli G, Ross C L, Moran        E, Salvatori M, Maghsoudlou P, Turmaine M, Delario G,        Al9Shraideh Y, Farooq U, Farney A C, Rogers J, Iskandar S S,        Burns A, Marini F C, De Coppi P, Stratta R J, Soker S. Discarded        human kidneys as a source of ECM scaffolds for kidney        regeneration technologies. Biomaterials 2013; 34:5915925.    -   23) Baiguera S, Macchiarini P, Ribatti D. Chorioallantoic        membrane for in vivo investigation of tissue-engineered        construct biocompatibily. J Biomed Mater Res 2012; 100: 1425e34    -   24) Totinelli G, Maghsaudlou P, Garriboli M, Riegler J, Orlando        G, Burns A J, Sebire N J, Smith V V, Fishman J M, Ghionzoli M,        Turmaine M, Birchall M A, Atala A, Soker S, Lythgoe M F,        Seifalian A, Pierro A, Eaton S, De Coppi P. A rat decellularized        small bowel scaffold that preserves villus crypt architecture        for intestinal regeneration. Biomaterials 2012; 33(12): 3401e10.    -   25) Navone S E, Marfia G, Invernici G, Cristini S, Nava S, Balbi        S, Sangiorgi S, Ciusani E, Bosutti A, Alessandri G, Slevin M,        Parati E A. Isolation and expansion of human and mouse brain        microvascular endothelial cells. Nat Protoc 2013; 8(9):1680993.    -   26) Salvatori M, Peloso A, Zambon J P, Patel T, Orlando G.        Extracellular Matrix Scaffold Technology for Bioartificial        Pancreas Engineering: State of the Art and Future Challenges. J        Diabetes Sci Technol 2014; 8(1):1599169.

Each of the references cited herein is incorporated by reference hereinin its entirety, or in relevant part, as would be apparent from thecontext of its citation.

From the disclosure herein it will be appreciated that, althoughspecific embodiments of the disclosure have been described herein forpurposes of illustration, various modifications may be made withoutdeviating from the spirit and scope of the disclosure.

What is claimed is:
 1. A method of regenerating a tissue or organ for asubject in need comprising: (a) providing an acellular tissue or organscaffold; (b) contacting the tissue or organ scaffold with a cellcharacteristic of the tissue or organ; (c) incubating the scaffold andcell ex vivo under static conditions; (d) implanting the incubatedscaffold and cell into the subject and establishing at least twovascular connections of the scaffold vasculature and the subjectvasculature; and (e) incubating the acellular scaffold until an intacttissue or organ is produced.
 2. The method of claim 1 wherein the cellis obtained from the subject.
 3. The method of claim 1 wherein thescaffold is contacted with at least 5×10⁷ cells.
 4. The method of claim1 wherein the cell is a vascular endothelial cell or an inducedpluripotent stem cell.
 5. The method of claim 1 further comprisingincubating ex vivo the scaffold and cell under ramping perfusionconditions.
 6. The method of claim 5 wherein the perfusion comprisesdelivery of at least 1×10⁸ cells at a rate spanning 2 ml/minute to 20ml/minute.
 7. The method of claim 1 further comprising attachment to thescaffold of an antibody product specifically binding to the cell.
 8. Themethod of claim 7 wherein the antibody product comprises an antigenbinding site for CD31.
 9. The method of claim 1 further comprisingattachment of an anti-thrombotic composition to the scaffold.
 10. Themethod of claim 9 wherein the anti-thrombotic composition is heparin.11. The method of claim 1 wherein the organ is a kidney, a pancreas or aliver.
 12. A method of decellularizing a tissue or organ comprising: (a)Perfusing an ex vivo tissue or organ with at least 50 volumes of adetergent; and (b) Rinsing the ex vivo tissue or organ with at least 50volumes of a neutral buffer.
 13. The method according to claim 12wherein the organ is selected from the group consisting of kidney,pancreas, and liver.
 14. The method according to claim 12 wherein theperfusing and rinsing are performed using at least two fluid inlets andat least one fluid outlet.
 15. The method according to claim 14 whereinthe flow rate of the two fluid inlets is about equal.
 16. The methodaccording to claim 12 further comprising an initial step of deliveringdistilled water to the organ or tissue at a rate of about 12 ml perminute for at least 10 hours.
 17. The method according to claim 12further comprising delivering at least one liter of 0.0025 w/w % DNasesolution to the detergent-treated tissue or organ prior to rinsing thetissue or organ.
 18. The method according to claim 12 wherein thedetergent is sodium dodecyl sulfate or Triton X-100.
 19. The methodaccording to claim 12 further comprising exposing the tissue or organ toa bactericidal agent.
 20. The method according to claim 19 wherein thebactericidal agent is betadine.
 21. The method according to claim 12wherein the neutral buffer contains 10 U/ml heparin.